Modulating Energy and Electron Transfer Processes in Photosystem II Of

Modulating Energy and Electron Transfer Processes in Photosystem II of

Chlamydomonas reinhardtii

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

Zoee Gokhale Perrine

Biophysics Graduate Program

The Ohio State University

2010

Dissertation Committee:

Dr. Ross Dalbey (Advisor)

Dr. Richard Sayre (Co-Advisor)

Dr. Terry Gustafson

Dr. Patrice Hamel

Zoee Gokhale Perrine

2010

Abstract

Photosystem II (PS II) is a water-plastoquinone oxidoreductase and is central to the process of oxygenic photosynthesis. It is associated with a peripheral light-harvesting complex (LHCII) or antenna that primarily functions in light absorption and transfer of excitation energy to the PS II reaction center (RC), where charge separation occurs.

Photosynthetic productivity is in part determined by the light utilization capacity of the peripheral antennae associated with PS II. Wild-type algae light saturate photosynthesis at 25% of full sunlight intensity. Hence under high light intensities, up to

~80% of absorbed photons may be dissipated as heat or fluorescence leading to a decrease in photosynthetic productivity. It was our objective to increase light utilization efficiency and productivities of algal cultures by optimizing the size of the PS II peripheral antenna. The lack of chlorophyll (Chl) b is known to preferentially decrease the size of the PS II peripheral antenna. Two approaches were adopted to modify Chl b accumulation and PS II antenna size. The first involved RNAi-mediated silencing of the

Chl b synthesis gene, CAO, which resulted in transgenics that had ~1.5-2.2 fold higher

Chl a/b ratios and reduced PS II peripheral antenna sizes compared to wild-type. This translated to a ~2 fold increase in light saturated rates of photosynthesis and 35% greater growth at high light intensities, while photoautotrophic growth at low light intensities was unimpaired. The second strategy involved the dynamic modulation of PS II antenna ii size through the post-transcriptional regulation of CAO by the light-responsive translational inhibitor protein, NAB1. Pigment analysis and Chl fluorescence induction using independent transformants expressing the modified CAO gene had increased Chl a/b ratios and reduced PS II antennae size under high light and the opposite phenotype under low light. The dynamic light-dependent modulation of PS II antenna size provided by this strategy is ideal for maximizing photon absorption and utilization during seasonal changes in light intensity.

The efficiency of forward electron transfer (ET) in PS II and hence photosynthetic productivity is also dependent, in large part, on the redox properties and energetic gaps between electron donors and acceptors which in turn can be influenced by the protein microenvironment of the redox cofactors in PS II. The monomeric ChlD1 on the active branch is the likely primary electron donor in PS II and the amino acid closest to it is D1-

T179. Our goal was to modify the local protein environment of ChlD1 to elucidate its function in charge separation and to study the effects of the protein environment on its redox properties. The resulting transgenics D1-T179S, D, H, N and I had impaired photoautotrophic growth and lower oxygen evolving activities in comparison to wild- type. The D1-T179N and I mutants also had elevated levels of basal PS II Chl fluorescence indicating an impairment in excitation energy transfer from the proximal antenna to the PS II RC. The combined results of the thermoluminescence and flash- induced Chl fluorescence decay measurements indicated that the mutagenesis of D1-

+ T179 to D, N or H induced a shift in the Em of P680/P680 to more positive values, causing an acceleration of charge recombination due to a decrease in the energetic gap between

iii

- - the primary radical pair and the S2QA (QB ) state. Our results show that the nature of the residue in position D1-179 is involved in mediating excitation energy transfer to the RC and is paramount in determining the redox properties of the primary electron donor in PS

II. The latter plays a critical role in balancing forward ET with charge recombination.

The primary electron acceptor in PS II is PheoD1 and its redox properties are modulated by the D1-E130 amino acid residue in Chlamydomonas which serves as a hydrogen bond donor to the PheoD1 head group. It has been shown that mutagenesis of

D1-130 to amino acids that weaken the hydrogen-bonding interaction to PheoD1 also shift

- the midpoint potential of the PheoD1/PheoD1 couple to more negative values, making forward ET transfer from PheoD1 to QA less probable. D2-Q129 is the inactive branch residue analogous to D1-E130 and it is found to be too distant from the PheoD2 head group to serve as a viable hydrogen bond donor. Our objective was to characterize the function of this highly conserved inactive branch residue by replacing it with a non- conservative leucine or a conservative histidine residue. The analysis of flash-induced

Chl fluorescence decay and thermoluminescence measurements indicated that the mutagenesis of D2-Q129 decreased the redox gap between QA and QB due to a lowering

- of the redox potential of QB. Further, the increased yield of S2QB charge recombination in the D2-Q129 mutants relative to wild-type also led to increased susceptibility to

3 photoinhibitory light presumably due to P680-mediated oxidative damage. This study provides insight into the extent of asymmetry between the two ET branches in PS II where analogous active (D1-E130L) and inactive (D2-Q129L) branch mutations can impact non-analogous cofactors of the two branches. The experimental data help to

iv increase our understanding of the specific impact of individual inactive branch amino acid residues on electron transfer in PS II.

Dedicated to my loving family

Acknowledgments

I would like to express my sincere gratitude to my research advisor, Dr. Richard

Sayre for his guidance, support and belief in me as a graduate student. Without his vision and scientific rigor, none of this would have been possible. I am also grateful to the members of my advisory committee, Dr. Ross Dalbey, Dr. Terry Gustafson and Dr.

Patrice Hamel for bearing with me through our inter-state interactions during the latter part of my PhD research and on one occasion, for listening to my research presentation while an unforgiving snow storm piled (what seemed like) 20 feet of snow outside.

I would like to acknowledge the assistance of Michal Sicner from Photon Systems

Instruments who provided valuable help on the workings of the kinetic fluorometer and thermoluminescence instruments.

Further I am grateful to have shared my laboratory experience with people who started out as colleagues and with time, became inseparable friends. I feel lucky to have been part of the professional scientific culture at the Donald Danforth Plant Science

Center in St. Louis, during the final two years of my PhD.

Finally, I feel indebted to my family for sharing in the trials and tribulations of my graduate school experience and for being my reticent punching bags when the going was tough. I will be eternally grateful to have succeeded in this challenging endeavor.

vii

Vita

2001...... B.Sc. Life Sciences and Biochemistry,

St. Xavier‘s College, Mumbai, India

2003...... M.Res. Bioinformatics,

University of Glasgow, Scotland, UK

2004 to present ...... Graduate Teaching and Research Associate,

The Ohio State University.

Publications

Gokhale, Z., and Sayre, R.T. (2009). Photosystem II, a Structural Perspective. In The

Chlamydomonas Sourcebook, D. Stern, ed (Boston: Academic Press), pp. 573-602.

Fields of Study

Major Field: Biophysics

viii

Table of Contents

Abstract...... ii

Dedication...... vi

Acknowledgments...... vii

Vita...... viii

List of Tables...... xiv

List of Figures ...... xvi

List of Abbreviations...... xxi

Chapter Page

1. Introduction...... 1

1.1 Overview of photosynthesis...... 1

1.2 Overview of photosynthetic light-harvesting antennae...... 4

1.2.1 Light-harvesting antenna of PS II...... 4

1.2.2 Light-harvesting antenna of PS I...... 12

1.2.3 Light-harvesting polypeptides...... 17

1.2.4 Regulation of light-harvesting capacity...... 21

1.2.4.1 Energy dependent quenching, qE...... 24

1.2.4.2 State transitions, qT...... 30

1.3 Overview of photosystem II...... 37

1.3.1 PS II crystal structure...... 38

1.3.2 D1 and D2 RC polypeptides...... 44

1.3.3 Primary electron donor, P680...... 46

1.3.4 Primary electron acceptor, PheoD1...... 52

1.3.5 Secondary electron acceptors, QA and QB...... 59

1.3.6 Oxygen evolving complex (OEC)...... 67

1.3.7 Chlz cycle...... 75

1.4 Implications of photosynthesis in biofuel production...... 77

1.5 This work...... 83

2. Effects of Modulating PS II Peripheral Antenna Size on Photosynthetic

Productivity and Growth of C. reinhardtii...... 85

2.1 Introduction...... 85

2.2 Materials and Methods...... 95

2.2.1 DNA constructs...... 95

2.2.2 Generation and screening of the CAO-RNAi transformants...... 100

2.2.3 Generation of NAB1 regulated CAO transformants...... 103

2.2.4 Chl fluorescence induction measurements...... 104

2.2.5 Non-denaturing polyacrylamide gel electrophoresis...... 105

2.2.6 Photoautotrophic growth of CC-424, CR-118, 133 and cbs-3...... 106

2.2.7 Light-saturation curves of CC-424, CR-118, 133 and cbs-3...... 106

2.2.8 Pigment determination by HPLC...... 107 x

2.3 Results...... 108

2.3.1 RNAi-mediated silencing of the CAO gene leads to transgenic algae

with altered Chl a/b ratios...... 108

2.3.2 CAO-RNAi transgenics have lower CAO mRNA levels...... 110

2.3.3 CAO-RNAi transgenics have truncated (intermediately-sized) PS II

antennae...... 112

2.3.4 Chl fluorescence rise kinetics are directly correlated with Chl b content and

size of the PS II antenna...... 114

2.3.5 Photosynthetic oxygen evolution of WT, CR-118, 133 and cbs-3...... 117

2.3.6 Photoautotrophic growth of WT, CR-118, 133 and cbs-3 cells...... 121

2.3.7 PCR confirmation of the NAB1 regulated CAO transgenics...... 124

2.3.8 Chl a/b ratios of the N1BS-CAO transgenics...... 127

2.3.9 Chl fluorescence induction in the N1BS-CAO transgenics...... 129

2.4 Discussion...... 131

3. Site-Directed Mutagenesis of D1-T179 Modifies the Redox Properties of the

Primary Electron Donor in Photosystem II...... 136

3.1 Introduction...... 136

3.2 Materials and Methods...... 143

3.2.1 Bioinformatics analysis...... 143

3.2.2 DNA constructs...... 144

3.2.3 C. reinhardtii chloroplast transformation...... 146

3.2.4 Growth of Chlamydomonas WT and D1-T179 mutants...... 147

3.2.5 Chl analysis...... 148

3.2.6 Assaying oxygen evolution...... 148 xi

3.2.7 Flash-induced Chl fluorescence measurements...... 149

3.2.8 Thermoluminescence measurements...... 150

3.3 Results...... 150

3.3.1 Protein environment of the monomeric Chls in type II RCs...... 150

3.3.2 Growth characteristics of site-directed mutants of D1-T179...... 153

3.3.3 Oxygen evolution measurements of WT and D1-T179 mutants...... 155

3.3.4 Basal Chl fluorescence (F0) and flash-induced Chl fluorescence relaxation

kinetics...... 157

3.3.5 Chl fluorescence decay in the presence of QB site inhibitors...... 161

3.3.6 Thermoluminescence properties of WT and D1-T179 mutants...... 164

3.4 Discussion...... 169

4. Site-Directed Mutagenesis of D2-Q129 Alters the Redox Potential of QB and

Charge Recombination in Photosystem II...... 181

4.1 Introduction...... 181

4.2 Materials and Methods...... 188

4.2.1 DNA constructs...... 188

4.2.2 C. reinhardtii chloroplast transformation...... 190

4.2.3 Growth of Chlamydomonas WT and mutant (D1-E130L, D2-Q129L,

D2-Q129H and D1-E130L/D2-Q129L) cells...... 191

4.2.4 Oxygen evolution measurements...... 191

4.2.5 Flash-induced Chl fluorescence induction and decay kinetics...... 191

4.2.6 Thermoluminescence measurements...... 192

4.2.7 Photoinhibition measurements...... 193

4.3 Results...... 193 xii

4.3.1 Photoautotrophic growth of WT and D2-Q129L, D2-Q129H,

D1-E130L and D1-E130L/D2-Q129L cells...... 193

4.3.2 Oxygen evolution activity...... 196

4.3.3 Flash-induced Chl fluorescence relaxation kinetics...... 198

4.3.4 Chl fluorescence decay in the presence of QB site inhibitors...... 203

4.3.5 Chl fluorescence induction kinetics of WT and D2-Q129L, D2-Q129H,

D1-E130L and D1-E130L/D2-Q129L cells...... 207

4.3.6 Thermoluminescence properties measured in the absence and presence of

20 µM DCMU...... 209

- 4.3.7 Effect of p-benzoquinone on S2QB charge recombination...... 217

4.3.8 Effects of photoinhibitory light treatment...... 220

4.4 Discussion...... 223

5. Summary...... 234

References...... 254

xiii

List of Tables

Table Page

1.1 LHCII proteins residues that are phosphorylated during state

transitions...... 32

1.2 Comparison of microalgae with other biodiesel feedstocks...... 82

2.1 Pigment composition of the PS II proximal and peripheral antenna proteins...... 89

2.2 List of primers used for cloning of the CAO-RNAi and

N1BS-CAO gene constructs...... 98

2.3 Primers used for real-time PCR analysis of the

CAO-RNAi (CR) transformants and CC-424 (WT)...... 102

2.4 Chl a/b ratios of independent CR transformants...... 109

3.1 List of primers used for site-directed mutagenesis of D1-T179...... 145

3.2 Position Specific Scoring Matrices (PSSMs) generated for

the Pfam00124 and CHL00003 protein families...... 152

3.3 Rates of steady-state oxygen evolution measured in whole

cells of the complemented WT and D1-T179 mutants...... 156

3.4 Chl fluorescence decay kinetics in the absence of 20 µM of DCMU...... 160

3.5 Chl fluorescence decay kinetics in the presence of 20 µM of DCMU...... 163

3.6 Possible coordination distances predicted through backbone-dependent

in silico mutagenesis of the D1-T179 residue...... 172

xiv

3.7 Free energy changes related to charge recombination in the

D1-T179 mutants...... 176

4.1 List of primers used for site-directed mutagenesis of amino acid residues

D1-E130 and D2-Q129...... 189

4.2 Steady-state oxygen evolution rates of WT, D2-Q129L, D2-Q129H,

D1-E130L and D1-E130L/D2-Q129L cells...... 197

4.3 Chl fluorescence decay kinetics of WT, D2-Q129L, D2-Q129H, D1-E130L

and D1-E130L/D2-Q129L cells in the absence of 20 µM DCMU...... 202

4.4 Chl fluorescence decay kinetics of WT, D2-Q129L, D2-Q129H, D1-E130L

and D1-E130L/D2-Q129L cells in the presence of 20 µM DCMU...... 206

4.5 TL peak temperatures for WT, D2-Q129L, D2-Q129H, D2-E130L and

D1-E130L/D2-Q129L...... 228

List of Figures

Figure Page

1.1 Schematic model of the major protein complexes involved in oxygenic

photosynthesis...... 3

1.2 Structural model of the intrinsic protein subunits within the PS II

supercomplex of higher plants at 24 Å...... 6

1.3 Three dimensional map of the PS II-LHCII supercomplex from

Chlamydomonas at 30 Å...... 8

1.4 Three dimensional structure of LHCII...... 11

1.5 The structural model of plant photosystem I at 3.4 Å resolution...... 13

1.6 Three-dimensional reconstruction of the Chlamydomonas PS I-LHCI

supercomplex at 30 Å resolution...... 16

1.7 Unrooted phylogenetics tree of Chlamydomonas LHCII proteins...... 20

1.8 Temporal sequence of photoacclimation responses...... 23

1.9 Schematic model for qE in plants...... 29

1.10 Proposed model for the migration of LHCII proteins during

transition from state 1 to state 2...... 35

1.11 View of the PS II complex...... 41

1.12 The organization of electron transfer cofactors in cyanobacterial

xvi

PS II reaction center crystals...... 43

1.13 PS II RC Chls and the surrounding protein environment...... 51

1.14 Protein environments of PheoD1 and PheoD2 in PS II...... 58

1.15 Energetics for electron transfer components in PS II based on the

redox potentials...... 60

1.16 View of the non-heme iron and the QA and QB binding pockets...... 66

1.17 The Mn4Ca cluster of the oxygen evolving complex of PS II...... 74

1.18 Photosynthetic biomass and biofuel production by green algae...... 79

2.1 Rates of photosynthesis and light absorption as a function of light intensity...... 87

2.2 Subunit structure of PS II–LHCII supercomplex...... 90

2.3 Scenario for light absorption and utilization by algae with large

and small PS II antennae at high light intensity...... 93

2.4 Gene constructs used for the modulation of Chl b synthesis in

Chlamydomonas...... 99

2.5 Real-time RT-PCR analysis of the CR transformants...... 111

2.6 Chl fluorescence induction in CC-424 (WT), CR transformants

and cbs-3...... 113

2.7 Relationship between Chl a/b ratio and % saturation...... 115

2.8 Visualization of the LHCII complex in CC-424 (WT), CR-118, 133

and cbs-3 via non-denaturing PAGE...... 116

2.9 Oxygen evolution rates of CC-424 (WT), CR-118, 133 and cbs-3 as a function of

light intensity and normalized based on Chl...... 119

2.10 Oxygen evolution rates of CC-424 (WT), CR-118, 133 and cbs-3

as a function of light intensity and normalized based on cell density...... 120

2.11 Photoautotrophic growth of the WT, CR-118 and133 and cbs-3 cells xvii

at low and high light intensities...... 123

2.12 PCR confirmation of the transgenics containing the NAB1 regulated

CAO gene constructs...... 126

2.13 Changes in Chl a/b ratios in the complemented WT (CAO-4, 22),

CC-2137 (WT), N1BSCAO and altN1BSCAO transgenics

during acclimation to low and high light...... 128

2.14 Changes in Chl fluorescence induction in the complemented WT (CAO-4, 22),

CC-2137 (WT), N1BSCAO and altN1BSCAO transgenics

during acclimation to low and high light...... 130

3.1 The protein environment of ChlD1 and the relative location of D1-T179...... 142

3.2 The growth characteristics of the WT and D1-T179 mutants

on TAP and High Salt media...... 154

3.3 Chl fluorescence decay curves measured in the

absence of 20 µM DCMU...... 159

3.4 Chl fluorescence decay curves measured in the

presence of 20 µM DCMU...... 162

3.5 Thermoluminescence characteristics of the WT and D1-T179

mutants measured in the absence 20 µM DCMU...... 166

3.6 Thermoluminescence characteristics of the WT and D1-T179

mutants measured in the presence of 20 µM DCMU...... 168

3.7 Scheme of charge recombination pathways in PS II...... 180

4.1 Structure of the acceptor side of Photosystem II...... 187

4.2 Growth characteristics of WT, D2-Q129L, D2-Q129H, D1-E130L

and D1-E130L/D2-Q129L (LL) cells...... 195

xviii

4.3 Flash-induced Chl fluorescence decay of WT, D2-Q129L, D2-Q129H, D1-

E130L and D1-E130L/D2-Q129L cells measured in the

absence of 20 µM DCMU...... 201

4.4 Chl fluorescence decay of WT, D2-Q129L, D2-Q129H, D1-E130L and D1-

E130L/D2-Q129L cells measured in the

presence of 20 µM DCMU...... 205

4.5 Chl a fluorescence induction transients of WT, D2-Q129L, D2-Q129H,

D1-E130L and D1-E130L/D2-Q129L cells...... 208

4.6 Thermoluminescence characteristics of the WT and D2-Q129 mutants

in the absence of 20 µM DCMU...... 211

4.7 Thermoluminescence characteristics of the D1-E130L and

D1-E130L/D2-Q129L mutants in the absence of 20 µM DCMU...... 212

4.8 Thermoluminescence characteristics of the WT and D2-Q129 mutants

in the presence of 20 µM DCMU...... 215

4.9 Thermoluminescence characteristics of WT, D1-E130L and D1-E130L/D2-

Q129L mutants in the presence of 20 µM DCMU...... 216

4.10 Thermoluminescence characteristics of WT and D2-Q129 mutants

after PBQ treatment...... 218

4.11 Thermoluminescence characteristics of the D1-E130L and D1-E130L/D2-Q129L

after PBQ treatment...... 219

4.12 Residual rates of oxygen evolution measured following photoinhibitory light

treatment of WT and D2-Q129 mutant thylakoids...... 221

4.13 Growth characteristics of the WT and D2-Q129 mutants under

high light (500 µmol light m-2 s-1)...... 222

4.14 Effect of in silico mutagenesis of the D2-Q129 residue...... 231 xix

5.1 Photosynthesis and biofuel production...... 236

5.2 Results summary of the NAB1-CAO transgenics

characterized in Chapter 2...... 240

5.3 Summary of important results of D1-T179 mutagenesis

described in Chapter 3...... 245

5.4 Results summary of Chapter 4...... 253

List of Abbreviations

ATP adenosine-5'-triphosphate

BChl bacteriochlorophyll

Chl chlorophyll

Cyt cytochrome

D1 and D2 photosystem II reaction center proteins

DCMU 3-(3,4-dichlorophenyl)-1,1-dimethylurea

DMBQ 2,6-dimethylbenzoquinone

ET electron transfer

F0 basal Chl fluorescence

Fv variable Chl fluorescence

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

HPLC High-performance liquid chromatography

HS High salt (photoautotrophic growth medium)

LHCI peripheral light-harvesting complex associated with photosystem I

LHCII peripheral light-harvesting complex associated with photosystem II

NADP nicotinamide adenine dinucleotide phosphate

NPQ non-photochemical quenching

P680 primary electron donor of photosystem II

PAGE polyacrylamide gel electrophoresis

PBQ p-benzoquinone

xxi

PCR polymerase chain reaction

Pheo pheophytin

PQ plastoquinone

PS I photosystem I

PS II photosystem II

QA primary quinone electron acceptor of photosystem II

QB primary quinone electron acceptor of photosystem I qE energy-dependent non-photochemical quenching qT state transitions

RC reaction center

TAP Tris-acetate-phosphate (photoheterotrophic growth medium)

TL thermoluminescence

xxii

Chapter 1

Introduction

1.1 Overview of photosynthesis

Photosynthesis is the process by which energy from sunlight is harvested and used by photosynthetic organisms to generate chemical energy for their cellular needs. Plants, green algae and cyanobacteria, carry out oxygenic photosynthesis in which light energy is used to generate NADPH and ATP required for the fixation of carbon dioxide to produce sugar. The overall biochemistry of photosynthesis for the formation of one glucose molecule may be written as:

6 CO2+ 12 H2O + Light  C6H12O6 + 6 O2 + 6 H2O

Oxygenic photosynthesis is absolutely vital in sustaining the biosphere as it simultaneously creates an aerobic environment as well as sources of reduced carbon. It can be described by two phases (i) a light-dependent phase (light reactions) during which

ATP and NADPH are formed and (ii) a light-independent phase (Calvin cycle) during which carbon dioxide (CO2) fixation and carbohydrate synthesis occurs. In photosynthetic eukaryotes, photosynthetic electron transfer and carbon fixation reactions are compartmentalized in the chloroplast. The light reactions of photosynthesis take place in the thylakoid membranes and involve the major thylakoid membrane protein 1 complexes as pictured in Fig. 1.1. Light is captured by the light-harvesting complexes

LHCI (dark green, labeled Lhca) and LHCII (lime green, labeled Lhcb) that are associated with photosystem I (PS I) and photosystem II (PS II) respectively. Linear electron transport is driven by the two photosystems and involves the transfer of electrons from water to NADP+, to form NADPH. Other components of the linear electron transport chain include plastoquinone (PQ), cytochrome b6f (Cytb6f), plastocyanin (PC), and PS I-bound ferredoxin (Fd). Reduced ferredoxin is used by a ferredoxin–NADP+ oxido-reductase to form NADPH. The linear electron transport pathway involving PS II and PS I causes protons to accumulate in the thylakoid lumen, leading to the buildup of a pH gradient. This ΔpH provides the proton motive force necessary for the chloroplast

ATP synthase to generate ATP. Under certain conditions cyclic electron transport around

PS I can be elicited. While linear electron transfer leads to the formation of ATP and

NADPH, cyclic electron transfer around PS I generates only ATP. Hence the balance of linear and cyclic electron transfer during oxygenic photosynthesis is an important regulator of intracellular ATP/NADPH ratios and downstream energy requiring processes

(Cardol et al., 2009). The products of the photosynthetic electron transfer, ATP and

NADPH, are consumed during CO2 fixation, where a stromal enzyme, RuBisCO, is responsible for catalyzing the incorporation of CO2 into Ribulose bisphosphate (RuBP), ultimately leading to the formation of sugars.

Fig. 1.1 Schematic model of the major protein complexes involved in oxygenic photosynthesis (http://www.photosynthesis.sbcs.qmul.ac.uk/nield/PS IIimages).

1.2 Overview of photosynthetic light-harvesting antennae

Photosystem I (PS I) and photosystem II (PS II) possess their own reaction centers

(RC) and peripherally located multi-subunit light-harvesting antenna systems. These antennae are responsible for capturing light energy and transferring it to the RCs where it is used to drive photochemistry. In plants and green algae, the light harvesting antennae for PS I and PS II are composed of the light-harvesting complexes I (LHCI) and II

(LHCII) respectively which bind chlorophyll a (Chl a), chlorophyll b, (Chl b) and carotenoids. LHCI and LHCII proteins have significant homology and are part of the light-harvesting complex (LHC) superfamily (Elrad and Grossman, 2004; Minagawa and

Takahashi, 2004). The primary role of the LHC proteins is in light-harvesting, although they also play a vital role in photoprotection of the photosynthetic apparatus under high light stress (Müller et al., 2001). This will be discussed in greater detail in later sections.

1.2.1 Light-harvesting antenna of PS II

The PS II-LHCII supercomplex is multi-subunit Chl-protein complex that resides in the thylakoid membranes of eukaryotic oxygenic photoautotrophs (reviewed in Dekker and Boekema, 2008; Minagawa, 2009; Gokhale and Sayre, 2009). The supercomplex collects and utilizes light energy to drive the energetically expensive transfer of water- derived electrons to the final PS II electron acceptor, plastoquinone. The D1 and D2 RC proteins form the very center of the supercomplex and house the redox active cofactors that participate in primary photochemistry. Flanking either side of the RC complex, are the Chl a containing proteins CP43 and CP47 which function to harvest and mediate

4 transfer of excitation energy from LHCII to the RC Chls. The D1, D2, CP43 and CP47 proteins form the RC core complex. At the periphery of the PS II core in plants and green algae is an additional light-harvesting antenna composed of LHCII proteins which account for up to 50% of the total Chl present in the thylakoid membranes. Hence, the light harvesting apparatus for PS II can be thought of as comprising of the core

(proximal) antenna made up of CP43 and CP47 and the peripheral (distal) antenna that is composed of LHCII. The peripheral antenna can further be classified as the major (outer) and more abundant trimeric antenna and the minor (inner) less abundant monomeric antenna (CP26 and CP29) (Minagawa and Takahashi, 2004) (Fig. 1.2, Nield et al.,

2000a).

Fig. 1.2 Structural model of the intrinsic protein subunits within the PS II supercomplex of higher plants at 24 Å (Nield et al., 2000a, Nat. Struct. Biol. 7: 44 – 47).

The three-dimensional structure of the Chlamydomonas PS II-LHCII supercomplex has also been obtained at 30 Å resolution using electron microscopy and single-particle analyses of negatively stained PS II complexes (Nield et al., 2000b). The PS II supercomplex particle isolated from C. reinhardtii was determined as being very similar to that from spinach suggesting a common basic unit consisting of two trimers of LHCII and two copies each of CP26 and CP29 per PS II core dimer (Fig. 1.3). The PS II supercomplex isolated from Chlamydomonas corresponded to the C2S2-type supercomplex (C and S refer to core complex and strongly associated LHCII trimers)

(Boekema et al., 1995; Dekker and Boekema, 2005). In plants however there can exists

C2S2M1 and C2S2M2 supercomplexes (M refers to moderately associated LHCII trimers).

Another monomeric LHCII antenna protein, CP24, is only present in plants and not green algae and hence is postulated to be required for binding M trimers to the core in higher plants.

Fig. 1.3 Three dimensional map of the PS II-LHCII supercomplex from Chlamydomonas at 30 Å (adapted from Nield et al., 2000b, J. Biol. Chem. 275: 27940 – 27946).

The structure of the major LHCII trimeric protein complex has been determined by

X-ray crystallography at 2.7 Å from spinach (Liu et al., 2004) and at 2.5 Å from pea

(Standfuss et al., 2005). Each LHCII monomer has three transmembrane helices (A, B and C) and two amphipathic helices (D and E) (Fig. 1.4). The two long, titled transmembrane helices A and B are related by a local near-two fold symmetry and are connected by a third shorter helix C (Kühlbrandt et al., 1994; Liu et al., 2004; Standfuss et al., 2005; reviewed in Barros and Kühlbrandt, 2009). The two short symmetry-related amphipathic helices on the lumenal side, helices D and E, link helix A to the C-terminus and helix B to helix C respectively (Fig. 1.4). Each LHCII monomer binds eight Chl a

(Chl 1-8), six Chl b (Chl 9-14), two lutein (L1-2) and one 9‘cis-neoxanthin molecules.

The Chl b, lipid and violaxanthin molecules are placed along the monomer-monomer interface. The centrally placed helices of each monomer, helix A and helix B are held together by a symmetrical pair of salt bridges, making this part of the complex structurally rigid (Kühlbrandt, 1994). The Arg and Glu residues that form these salt bridges are conserved throughout the LHC gene family (Barros and Kühlbrandt, 2009).

The central structural motif of the LHCII protein is highly conserved throughout the LHC superfamily and includes the helices A and B, the two lutein molecules which are placed in the grooves at either side of the helix pair, and the 3 pairs of symmetry-related Chl a molecules (Chl 1/Chl 4, Chl2/Chl5, Chl 3/Chl 6). The amino acid side chains or backbone carbonyls that serve as ligands to the central Mg atoms of the bound Chls have been determined and are conserved. Chls 1–5, 8, 12 and 13 are coordinated by conserved amino acid side chains where as Chls 6, 9, 11 and 14 are coordinated by main chain

9 carbonyls via a water molecule. The central Mg of Chls 7 and 10 do not have protein ligands but are coordinated by the head group of phosphatidylglycerol and the Chl 13 oxygen respectively. In the LHCII protein, site specificity for binding Chl a or Chl b is determined by whether a hydrogen bonding partner for the Chl b formyl group is present.

In vitro reconstitution studies have shown that Chl a sites can bind Chl b and vice versa

(Giuffra et al., 1996; Pagano et al., 1998). However, there is no X-ray crystallographic evidence for mixed occupancy in vivo and each binding site seems specific for its corresponding pigment. The hydrogen bond donors required for proper binding of Chl b are largely conserved in the LHC proteins. Hence a minimum set of Chls that are common to most LHCs include Chls a 1–6 and 8, and Chls b 12 and 13. This is broadly consistent with the experimentally determined Chl content and a/b ratios of the minor

LHCs where CP29 binds six Chl a and two Chl b molecules (Sandonà et al., 1998) and

CP26 binds six Chl a and three Chl b molecules (Croce et al., 2002).

Fig. 1.4 Three dimensional structure of LHCII. Cyan, Chl a; green, Chl b; dark orange, lutein; light orange, 9‘cis-neoxanthin; pink, lipid; yellow, violaxanthin (Liu et al., 2004,

Nature 428: 287 – 292; Barros and Kühlbrandt, 2009, Biochim. Biophys. Acta 1787: 753

– 772).

1.2.2 Light-harvesting antenna of PS I

The core of the PS I-LHCI supercomplex is formed by the two principal PS I RC subunits, PsaA and PsaB that carry out electron transfer from reduced plastocyanin to ferredoxin. Two additional RC proteins are found exclusively in plants and green algae,

PsaG and PsaH, which provide contact sites for the binding of peripheral LHCI (Ben-

Shem et al., 2003; Amunts et al., 2007) and LHCII (Lunde et al., 2000) antennae respectively (Fig. 1.5). All eukaryotic PS I cores characterized thus far are monomers

(Amunts et al., 2007) while trimeric PS I cores have been observed in cyanobacteria

(Jordan et al., 2001). With the exception of a few Chls that take part in photochemistry, majority of the Chls bound by the core complex of PS I function in light-harvesting. In plants and algae, the LHCI peripheral antennae proteins are shown to be asymmetrically bound to the PsaG/F/K side of the PS I core, with stronger binding at the G-pole (PsaG end) rather than the K-pole (PsaK end) (Germano et al., 2002; Kargul et al., 2003;

Amunts et al., 2007). It is evident from the 3.4 Å crystal structure of the PS I-LHCI supercomplex from pea, that the peripheral LHCI antenna is half-moon shaped (―LHCI belt‖) and is formed by four LHCI proteins, Lhca1, Lhca4, Lhca2 and Lhca3, starting from the G-pole (Amunts et al., 2007). The half-moon shape of the peripheral LHCI antenna and increased flexibility among its monomers is thought to aid in light harvesting and excitation energy migration in changing light environments (Amunts et al., 2007).

Fig. 1.5 The structural model of plant photosystem I at 3.4 Å resolution (Amunts et al.,

2007, Nature 447: 58 – 63).

Unlike the LHCII proteins which have been shown to form monomers or trimers, the

LHCI proteins assemble into dimers. In Chlamydomonas, the peripheral LHCI antennae are divided into two subcomplexes, LHCI-705 and LHCI-680, so called due to their characteristic 77 K fluorescence peaks (Bassi et al., 1992). Although the algal and plant

PS I-LHCI supercomplexes share similar architectures, one of the key differences between the two is the number of bound LHCI polypeptides per PS I core. The 4.4 Å and

3.4 Å pea PS I-LHCI crystal structures show the presence of only 4 LHCI monomers attached to the PS I core (Ben-Shem et al., 2003; Amunts et al., 2007) whereas biochemical analyses have suggested that four (Croce et al., 2002; Klimmek et al., 2005) or eight (Boekema et al., 2001) LHCI proteins may bound to the PS I core in higher plants. Single particle analysis of the PS I-LHCI supercomplex from Chlamydomonas have suggested that up to 11 (Kargul et al., 2003) or 14 (Germano et al., 2002) LHCI polypeptides are found to bind to the PS I core (Fig. 1.6). Furthermore, while 168 Chls were assigned to the PS I-LHCI supercomplex in the 3.4 Å pea crystal structure (Amunts et al., 2007), 215 Chls were estimated for the PS I-LHCI supercomplex isolated from

Chlamydomonas (Kargul et al., 2003). By assuming that 100 Chls bind to the PS I core complex in both structures, the estimated number of Chls in the Chlamydomonas LHCI antenna is much higher than the number of Chls in the plant LHCI antenna. Hence, the light harvesting apparatus for PS I is possibly larger in green algae than in higher plants and it is thought that two ―LHCI belts‖ might exist in algae rather than one as in higher plants (Kargul et al., 2003).

While the structure of the four LHCI monomers maintain the previously described

LHCII fold of two long, tilted, transmembrane helices (analogous to helices A and B in

LHCII) and a shorter third one (corresponding to helix C), the conservation of the fourth helix (D) is not apparent (Amunts et al., 2007). In addition to binding 14 Chls, LHCI proteins also bind ―gap Chls‖ in the region between LHCI and the core complex. These pigments are thought to facilitate the connectivity between the LHCI proteins and the PS

I core (Melkozernov et al., 2006).

Fig. 1.6 Three-dimensional reconstruction of the Chlamydomonas PS I-LHCI supercomplex at 30 Å resolution (Kargul et al., 2003, J. Biol. Chem. 278: 16135 –

16141).

1.2.3 Light-harvesting polypeptides

The genome sequence of Chlamydomonas predicts 9 LHCA and 12 LHCB genes that encode the LHC proteins associated with PS I and PS II respectively (Minagawa and

Takahashi, 2004; Elrad and Grossman, 2004). The major LHCII proteins that form the outer trimeric antenna are encoded by nine genes LHCBM1—9 (M refers to major).

Based on their phylogenetic relationships, the major LHCII proteins can be classified under four distinct types as shown in Fig 1.7. The genes in the type I class share 89-99% sequence identity and include 5 members LHCBM3, LHCBM4, LHCBM6, LHCBM8 and

LHCBM9 (Minagawa, 2009). Type III genes share 99% similarity and include LHCBM2 and LHCBM7. Type II (LHCBM5) and IV (LHCBM1) each have one member as shown

(Fig. 1.7). As expected the Chl binding protein ligands in the polypeptides encoded by

LHCBM1-9 are conserved within the LHC family (Standfuss et al., 2005; Barros and

Kühlbrandt, 2009). The structural motif required for trimerization of LHCII proteins in vitro (WYXXXR) (Hobe et al., 1995) is completely conserved in the major LHC proteins type III and IV but only partially so in type I and II in Chlamydomonas. This is unlike the case in higher plants where all classes of major LHCII proteins have the trimerization motif (Hobe et al., 1995; Minagawa, 2009). Two-dimensional protein mapping of light- harvesting proteins associated with PS II in Chlamydomonas, has revealed that LHCII type I, III, and IV proteins are the predominant components of the major trimeric antenna

(Stauber et al., 2003). However, in a surprising result, the type II LHCII protein,

LHCBM5, was identified in protein spots excised from two-dimensional maps of PS I.

LHCBM5 was later detected as a mobile component of the major LHCII antenna during state transitions (Takahashi et al., 2006).

The two minor LHCII proteins, CP29 and CP26, are conserved throughout

Chlorophyta, however, an additional minor antenna protein, CP24, found in plants, has not been observed in green algae (Minagawa and Takahashi, 2004). A transiently expressed antenna protein, LHCB7, which is suggested to have a specialized function during plastid development in plants (Klimmek et al., 2006) and shares 31 and 46% identity to CP29 and CP26 respectively, is found to be conserved in several green algae

(Minagawa and Takahashi, 2004). CP26 and LHCB7 contain trimerization motifs, whereas CP29 does not (Hobe et al., 1995; Minagawa, 2009). The CP26 protein has been shown to undergo trimerization in an Arabidopsis mutant deficient in two of the major

LHCII proteins, Lhcb1 and 2 (Ruban et al., 2003). Recently it was demonstrated that the two monomeric LHCII proteins (CP26 and CP29) and the type II major LHCII protein

(LHCBM5) were found to be associated with the PS I-LHCI supercomplex during state transitions to state 2 (Takahashi et al., 2006). Like the major trimeric LHCII proteins, the

CP26, CP29 and LHCBM5 proteins are phosphorylated upon transition to state 2.

Interestingly, of the LHCII proteins, these three are also most similar to the LHCI proteins. These observations have led to the proposition that the CP26, CP29 and

LHCBM5 proteins that were previously thought to localize only to the PS II complex, shuttle between PS I and PS II during state transitions and act as docking sites for the trimeric LHCII proteins in both PS I and PS II (Takahashi et al., 2006).

The light harvesting antenna for PS I in Chlamydomonas is formed by the gene products of nine genes LHCI1-9. Among them LHCA1 and LHCA3 are most abundant

(Hippler et al., 2001) and are conserved in higher plants. Features such as a very short sequence between helices C and A or a six-residue insertion at the beginning of helix B that help classify them as either Type I or Type III respectively (Tokutsu et al., 2004).

However with the exception of LHCAI and LHCA3 the other LHCI proteins are not well conserved in algae compared to plants and are more divergent when compared to LHCII proteins (Elrad and Grossman, 2004). By analogy to the minor LHCII proteins which lie between the major LHCII and PS II RC core, it is thought that LHCI type I and III proteins function as the inner antennae for PS I and the five other Chlamydomonas LHCI polypeptides (LHCI 2, 4, 5, 6, 7, 8 and 9) function as distal antenna to the PS I core

(Tokutsu et al., 2004). In comparing the subunit compositions of the PS I-LHCI supercomplex and LHCI by proteomic analysis, it was observed that while the wild-type

PS I-LHCI supercomplex contained all of the nine LHCA polypeptides, the LHCI complex retained only six LHCA polypeptides, where LHCA3 and two minor (lower abundance) polypeptides, LHCA2 and LHCA9, were lost during the purification procedure. Hence it was suggested that LHCA2, LHCA3, and LHCA9 are not required for the formation of the oligomeric structure of LHCI and that the association of these proteins in the LHCI complex is stabilized by the presence of the PS I core (Takahashi et al., 2004).

Fig. 1.7 Unrooted phylogenetics tree of Chlamydomonas LHCII proteins (Minagawa,

2009, In Chlamydomonas Sourcebook, D. Stern, ed (Boston: Academic Press), pp. 503 –

539).

1.2.4 Regulation of light-harvesting capacity

As its name suggests, the primary function of the light harvesting complex is to make the photosynthetic process more efficient by collecting solar energy and transmitting it to the RCs where photochemistry takes place. In their natural environmental, photosynthetic organisms have to deal with frequent fluctuations in light quantity and quality and often have large antennae to maximize the efficiency of light harvesting at limiting light intensities. However, having a large antenna proves detrimental to the organism under high light intensities due to saturation of electron transfer and over-absorption of photons.

In wild-type algae, which have large PS II antennae, photosynthesis is saturated at only

25% of full sunlight intensity. However the rate of photon absorption does not saturate.

This results in the formation of Chl excited states that cannot be utilized for photosynthesis. Under these conditions the increased lifetime of singlet excited Chl increases the propensity of triplet Chl formation through intersystem crossing which can transfer its excitation energy to the triplet ground state of oxygen resulting in singlet oxygen generation (Müller et al., 2001). Further, charge recombination reactions between

+ P680 and the acceptor side of PS II can also lead to the formation of triplet Chl and reactive oxygen species (ROS) production (Krieger-Lizkay et al., 2008). Hence photosynthetic organisms have developed short and long term responses that enable them to acclimatize to the prevailing light environment thereby reducing the deleterious effects of excess excitation and minimizing ROS-induced photodamage. These responses include

1) energy dependent non-photochemical quenching (qE-quenching) that involves the thermal dissipation of excess excitation energy, 2) adjustment of absorption cross-

21 sections of PS I and PS II by the redistribution of mobile LHCII antenna proteins (state transitions, qT), and 3) the transcriptional and translational control of LHC gene expression to control antenna size which will be discussed further in Chapter 2 (Teramoto et al., 2002; Durnford et al., 2003; Mussgnug et al., 2005).

The first two, qE and qT, are considered short term photoacclimation responses and are measurable components of non-photochemical quenching of chlorophyll fluorescence

(NPQ) while the third is a longer term photoacclimation response (Eberhard et al., 2008,

Fig. 1.8). The activation of NPQ photoprotective processes under conditions of excess light can lead to the dissipation of up to ~80% of absorbed photons and results in decreases in overall photosynthetic productivity (Polle et al., 2002). It is known that decreasing the size of the PS II antennae can alleviate light saturation rates of photosynthesis and increase culture productivities under high light. In Chapter 2, we describe genetic strategies to modulate PS II antennae size for the maximization of photon capture and utilization under all light intensities to increase year-round algal culture productivities.

Fig. 1.8 Temporal sequence of photoacclimation responses (Eberhard et al., 2008, Annu.

Rev. Genet. 42: 463 – 515).

1.2.4.1 Energy dependent quenching, qE

Energy dependent quenching, qE, is induced by acidic thylakoid lumen pH which occurs during high light exposure (reviewed in Horton et al., 1996; Müller et al., 2001).

The decrease in lumenal pH causes the protonation of PS II proteins involved in qE and the activation of violaxanthin de-epoxidase (VDE) that is responsible for the conversion of violaxanthin to zeaxanthin via antheraxanthin as part of the xanthophyll cycle (Eskling et al., 1997; Niyogi et al., 2005; Fig. 1.9). The induction of qE by high light, in dark adapted Chlamydomonas cells, displays biphasic kinetics (Niyogi et al., 1997; Niyogi,

2009). The first and rapid phase of qE occurs within a few seconds and depends on the generation of a pH gradient, the presence of lutein and low levels of zeaxanthin and antheraxanthin. The second and slower phase occurs on the order of minutes and is correlated with the formation of zeaxanthin and antheraxanthin resulting from VDE activity (Niyogi et al., 1997). Both phases are however dependent on the generation of a pH gradient. Insertional mutants of Chlamydomonas, npq1 and npq2, that are defective in

VDE activity and the zeaxanthin epoxidase (ZEP1) gene respectively show alterations in

NPQ characteristics (Niyogi et al., 1997). The second phase of qE is specifically affected in the npq1 mutant, which manifests as a 25% reduction of qE. The npq2 mutant that is defective in ZEP1, the enzyme that converts zeaxanthin and antheraxanthin to violaxanthin under low light, manifests a sustained form of NPQ due to permanently increased levels of zeaxanthin, which does not increase further upon high light exposure

(Niyogi et al., 1997). Additionally, the lack of lutein as in the lor1 mutant and in the npq1 lor1 double mutant causes a 50% and 90% reduction in qE respectively, suggesting that

24 zeaxanthin, antheraxanthin as well as lutein contribute to qE in Chlamydomonas (Niyogi et al., 1997).

NPQ mutants that are deficient in qE but have fully functional xanthophyll cycles have been isolated in Chlamydomonas and can help identify PS II proteins that play a role in qE induction. One such mutant is npq5, which was found to have a normal xanthophyll cycle and state transitions, but manifested a 90% decrease in qE (Elrad et al.,

2002). The mutation was revealed as being in the LHCBM1 gene which encodes a protein that is part of the trimeric LHCII antenna (Elrad and Grossman, 2004; Minagawa and

Takahashi, 2004). Studies involving the Arabidopsis npq4 mutant, in which the xanthophyll cycle is fully functional, revealed that the PS II subunit PsbS is a vital component of qE (Li et al., 2000) and that it is protonated at the onset of qE (Li et al.,

2004). Although the PsbS gene is present in the Chlamydomonas genome, the PsbS protein has not been detected in Chlamydomonas (Allmer et al., 2006). More recently the npq4 mutant of Chlamydomonas was characterized that had a fully operational xanthophyll cycle at high light but yet was unable to generate qE (Peers et al., 2009).

This mutant was shown to be deficient in two of the three genes encoding LHCSR

(L1818), an ancient member of the light harvesting complex superfamily. The LHCSR protein was suggested as being the protein with a function analogous to that of PsbS in plants that helps in switching the antenna of PS II between its light-harvesting and photoprotective functions (Peers et al., 2009).

Although NPQ has been phenomenologically documented for a long time, the physical basis of feedback de-excitation or energy-dependent quenching (qE) has

25 remained elusive until recently. As previously mentioned, qE involves the thermal dissipation of excess energy of Chl singlet excited states (1Chl*) in PS II of green plants and algae under high light, so as to minimize the production of reactive oxygen intermediates. In excess light, a low thylakoid lumen pH activates formation of the carotenoid zeaxanthin from violaxanthin via the xanthophyll cycle and drives protonation of a PS II subunit, PsbS, in plants that is necessary for qE (Li et al., 2000; Li et al., 2004).

It has been shown through transient absorption (TA) kinetics of spinach thylakoids excited at 664 or 683 nm and probed in the spectral region from 530 to 580 nm, that there is direct carotenoid excitation during qE (Ma et al., 2003). The kinetics in this spectral region are characteristic of the S1  Sn transition of xanthophyll and were found to be significantly different under the quenched (maximum NPQ) and unquenched (no NPQ) conditions. Further, ultrafast TA spectroscopy in the near infrared region (1000 nm) where the carotenoid radical cation species (Car•+) is known to exhibit strong absorption, showed a qE-dependent transient evolution of a Car•+ radical cation (Holt et al., 2005).

Subtraction of the quenched trace from the unquenched trace and a corresponding fit of the data produced a single exponential rise and decay of ~11 ps and ~150 ps, respectively. The rise component is characteristic of qE dependent quenching due to the transient formation of Car•+. Consistent with this observation, the near infrared TA kinetics from isolated thylakoids of the npq1 and npq4 mutants of A. thaliana specifically impaired in the qE component of NPQ, did not show transient Car•+ formation. Further, transient Car•+ signals in npq2 and npq2lut2 (a mutant that constitutively accumulates zeaxanthin but also lacks lutein) were very similar to each other providing evidence that

26 the Car•+ transiently formed in the wildtype thylakoids is a zeaxanthin radical cation species (Holt et al., 2005). Hence, based on the near infrared TA data the proposed model for the physical mechanism for qE involved the participation of three species including

(i) a bulk Chl pool (Chlbulk), (ii) a Chl-Zeaxanthin (Zea) heterodimer that quenches

–• excited Chlbulk molecules, and (iii) a charge-separated ground state consisting of Chl and

Zea+•. The charge separated state is formed from the relaxation of the (Chl-Zea)* excited state. According to the model, the ~11-ps rise component was attributed to the transfer of excitation energy from the Chlbulk molecules to Chl-Zea heterodimer which undergoes a fast relaxation to the charge separated ground state in < 1ps ensuring efficient quenching and preventing the back transfer of excitation energy to Chlbulk. The ~150 ps kinetic decay component was attributed to the loss of the Zea+• signal due to charge recombination with Chl-• (Holt et al., 2005). More evidence for the involvement of the zeaxanthin as the specific carotenoid species involved in qE quenching was obtained from ultrafast near infrared TA kinetics on isolated major and minor LHCII complexes of the wild-type and npq2 mutant of Arabidopsis (Avenson et al., 2008). The npq2

Arabidopsis mutant is deficient in zeaxanthin epoxidase and constitutively accumulates zeaxanthin (Niyogi et al., 1998). Hence LHCII proteins isolated from dark-adapted wild- type and the npq2 mutant were shown to be specifically enriched in violaxanthin and zeaxanthin, respectively. Since the kinetic profile at 1000 nm of wild-type and npq2 major LHCII proteins as well as wild-type minor LHCII proteins, had bi-exponential decays and lacked any rise components, the observed kinetic changes were attributed solely due to chlorophyll excited state absorbance dynamics, rather than transient Car•+

27 formation (Avenson et al., 2008). In contrast, the 1000 nm TA kinetics of the monomeric

(minor) LHCII proteins isolated from the npq2 mutant that were enriched in zeaxanthin, exhibited a small rise component with a time constant of 2.9 ps, followed by bi- exponential decay. The near infrared difference profile obtained from the monomeric

LHCII proteins of the wild-type and npq2 mutant was characterized by 5.2 ps and 238 ps rise and decay components respectively and were comparable to those of the corresponding rise and decay time constants of the difference TA kinetics acquired using isolated thylakoids from wild-type (Holt et al., 2005). Hence this provided evidence for the involvement of carotenoid radical cation generation in isolated minor light-harvesting complexes that bind zeaxanthin, in charge transfer quenching during qE (Avenson et al.,

2008). However, it was estimated that only 0.5% of isolated minor LHC complexes participated in charge transfer quenching in vitro, whereas the contribution in isolated thylakoids was > 80 times higher. It was concluded that minor LHCII complexes which bind zeaxanthin are the sites of charge transfer quenching in vivo, however the extent of quenching is modulated by the transmembrane pH gradient, the PsbS protein (in plants), and other protein-protein interactions. The physical mechanism for qE dependent quenching has not as yet been characterized in Chlamydomonas (Niyogi, 2009).

Fig. 1.9 Schematic model for qE in plants (adapted from Niyogi et al., 2005, J. Exp. Bot.

56: 375 – 382).

1.2.4.2 State transitions, qT

The second component of NPQ is the quenching of PS II Chl fluorescence via a process termed state transition, during which the relative excitation of PS I and PS II is modulated by the migration of mobile LHCII proteins between the two photosystems.

During state transitions, state I and state II are induced by the preferential excitation of

PS I (700 nm) and PS II (650 nm) respectively (reviewed in Wollman, 2001). It is thought that illumination conditions that lead to the preferential excitation of PS II lead to the reduction of the plastoquinone (PQ) pool and the docking of plastoquinol (PQH2) to the Q0 site of the cyt b6f complex. This activates an LHCII kinase which phosphorylates specific LHCII proteins (shown in Table 1.1) The mobile portion of the LHCII complex is then displaced from PS II and becomes associated with PS I, thus redistributing excitation energy between the two photosystems (reviewed in Rochaix, 2009).

Thylakoids in state 1 and state 2 can be distinguished based on Chl fluorescence emission at 77K where state 1 is characterized by a higher ratio of F685 and F695 (PS II Chl fluorescence emission) to F720 (PS I Chl fluorescence emission) emission and state 2 is characterized by a lower ratio of F685 and F695 to F720 (Depège et al., 2003; Takahashi et al., 2006). Additionally, a lower quantum yield for PS I photochemistry is apparent in

State 1 whereas a lower quantum yield for PS II and a higher quantum yield of PS I reactions is observed in State 2. The reversible phosphorylation of the mobile (major and minor) LHCII proteins is necessary for their migration to PS I during the transition from state I to state II. The phosphorylation sites in the LHCII proteins have been determined and are found to be clustered at the interface between the PS II core and LHCII antennae

(Turkina et al., 2006) (Table 1.1). Studies involving the stt7 Chlamydomonas and STN7

Arabidopsis state transition mutants have clearly demonstrated the involvement of STT7 and its Arabidopsis homolog, STN7, in LHCII phosphorylation and state transition

(Depège et al., 2003; Bellafiore et al., 2005). STT7 and STN7 encode thylakoid associated Ser-Thr protein kinases that are activated during state transition from state 1 to state 2. However it is not clear whether LHCII proteins are the immediate substrates of

STT7 and STN7 or whether they are part of a signal cascade that involves another downstream kinase.

Protein Residue Phosphorylated Under State 2

* indicates phosphorylated under State 1 and State 2 CP26 T10 Minor LHCII CP29 T7*, T11, T17, T18, T20, T33*, S103

LHCBM1 T27

LHCBM4 T19, T23

Major LHCII LHCBM6 T18, T22

LHCBM8 T19, T23

LHCBM9 T19, T23

Based on results from Turkina et al., 2006

Table 1.1 LHCII proteins residues that are phosphorylated during state transitions.

There is overwhelming evidence that supports the association of LHCII with the PS I complex during state 2. Crosslinking studies of PS I preparations from Arabidopsis, using dithio-bis(succinimidylpropionate), have shown that two to three fold more LHCII is associated with PS I in state 2 than in state 1 and that the PS I-H,-L, and -I subunits contain the docking sites for binding the mobile LHCII antenna (Zhang et al., 2004).

Further while LHCB1 was found associated with PS I in plants lacking the PS I-K subunit, PS I from the PS I-H, -L, or -O mutants contained only about 30% of LHCB1 compared with the wild type suggesting the importance of these PS I subunits in binding the mobile LHCII antenna. The involvement of the PS I-H subunit in binding LHCII antennae during state transitions is consistent with the finding that Arabidopsis mutants lacking PsaH are deficient in state transitions (Lunde et al., 2000). More recent studies have shown that PS I-LHCI supercomplexes isolated from Chlamydomonas reinhardtii cells locked in state 1 or state 2 have three LHCII proteins associated with the PS I–LHCI supercomplex only in state 2 (Takahashi et al., 2006). These three LHCII proteins were found to be phophorylated in state 2 (Turkina et al., 2006) and were identified as the two minor monomeric LHCII proteins (CP26 and CP29) and one major type II LHCII protein

(LHCBM5). Interestingly, these three LHCII proteins were most similar in sequence to the LHCI proteins. These observations provided evidence that the CP26, CP29, and

LHCBM5 proteins that were previously believed to be solely associated with PS II, shuttle between PS I and PS II during state transitions, and act as docking sites for the trimeric LHCII proteins in both PS I and PS II (Takahashi et al., 2006) (Fig. 1.10).

Analysis of Chlamydomonas RNAi mutants of the CP29 and CP26 minor LHCII

33 monomeric proteins demonstrated that CP29 is crucial for the formation of the PS

I/LHCI/LHCII supercomplex during state transitions whereas CP26 is not, although both are found to be tightly bound to PS I in state 2 (Tokutsu et al., 2009). Another important result that emerged from these mutants was that while CP29 is required for the re- association of the mobile LHCII with PS I during state transition to state 2, it is not essential for the dissociation of mobile LHCIIs from PS II. Hence other phosphoryated

PS II proteins such as CP26, CP43 and D2 are thought as being sufficient to undock mobile LHCII antennas from PS II during state transitions (Tokutsu et al., 2009).

Fig. 1.10 Proposed model for the migration of LHCII proteins during transition from state

1 to state 2 (Takahashi et al., 2006, Proc. Natl. Acad. Sci. U.S.A. 103: 477 – 482).

In Chlamydomonas 80% of LHCII is involved in state transitions compared to 15-

20% in Arabidopsis (Delosme et al., 1996). Hence, it is thought that that state transitions fulfill another function apart from the re-distribution of excitation energy between PS I and PS II. It has been demonstrated that state transitions play a role in maintaining the levels of intracellular ATP (Bulté et al., 1990) and more recent data involving single and double mutants defective for state transitions and/or mitochondrial respiration have shown that state transitions provide important energetic contributions in the form of ATP that are necessary for efficient carbon assimilation (Cardol et al., 2009). Hence it is postulated that state transitions in Chlamydomonas play a significant physiological role in meeting cellular energy demands which is achieved by modulating the relative excitation of PS II and PS I. Linear electron flow between PS II and PS I generates ATP and

NADPH, whereas cyclic electron transfer that occurs upon the preferential excitation of

PS I (state 2), generates only ATP. Hence, an imbalance in ATP/NADPH ratios caused by a decrease in intracellular ATP levels may be rectified by the activation of state transitions to state 2 and cyclic electron transfer.

1.3 Overview of photosystem II

Like the photosystem II peripheral antenna, the components of the photosystem II reaction center (PS IIRC) complex have an essential role in determining the efficiency of electron transfer and photosynthetic productivity. During the last decade substantial progress has been made in the elucidation of the structure and function of the PS II apparatus. Much of this progress has been achieved using either prokaryotic cyanobacterial PS II complexes, Chlamydomonas chloroplast PS II complexes, or the detergent-fractionated thylakoid membranes of flowering plants such as spinach.

Undoubtedly, the crystal structure from the thermophilic cyanobacteria has had the greatest impact on our understanding of structure–function relationships in PS II (Zouni et al., 2001; Kamiya and Shen, 2003; Ferreira et al., 2004; Loll et al., 2005; Guskov et al.,

2009). This structure has also served as a model for the chloroplast PS II complex, since the geometry of the RC chromophores seem precisely conserved although some differences are evident in the protein scaffolding at the stromal ends of the transmembrane (TM) helices (Buchel and Kuhlbrandt, 2005). Several reviews have been published on PS II structure and function, covering early events in primary charge separation (Diner and Rappaport, 2002; Vassiliev and Bruce., 2008; Rappaport and

Diner., 2008), structural studies (Ruffle and Sayre, 1998; Minagawa and Takahashi,

2004; Kern and Renger, 2007; Gokhale and Sayre, 2009), and the mechanism of water oxidation (Nelson and Yocum, 2006; Renger, 2007; Wydrzynski, 2008; Barber, 2008).

1.3.1 PS II crystal structure

PS II is a multi-subunit protein–cofactor complex embedded in the thylakoid membrane, which catalyzes the oxidation of water and the reduction of plastoquinone.

The holocomplex consists of no less than 20 protein subunits and has at least 77 cofactors including 35 Chl a, 12 carotenoid and 25 integral lipid molecules. X-ray diffraction studies have led to the elucidation of cofactor–protein interactions at near-atomic resolution (Zouni et al., 2001; Kamiya and Shen, 2003; Ferreira et al., 2004; Biesiadka et al., 2004; Loll et al., 2005; Guskov et al., 2009). Substantial knowledge regarding the identity and function of the redox active components had previously been generated using biophysical approaches (reviewed in Diner and Rappaport, 2002; Rappaport and Diner,

2008). Significantly, the predictions inferred from biophysical investigations have by and large been consistent with the structural information obtained from PS II crystals. The initial PS II-specific events leading to the generation of a charge-separated state involve excitation energy transfer from the CP43 and CP47 proteins of the proximal antenna complexes to the chlorins of the PS II RC. Energy transfer leads to the excitation of the

* primary electron donor of PS II, P680, leading to the formation of P680 . Subsequent charge transfer results in the reduction of the primary electron acceptor, a pheophytin a molecule (PheoD1) in a few picoseconds. The reduced primary acceptor then transfers an electron to the first stable (long-lived) electron acceptor, a bound plastoquinone (QA) molecule. The rate-limiting step of PS II ET is electron flow from QA to the plastoquinone QB bound at the QB-binding site. Following two rounds of reduction and subsequent protonation, the plastoquinol formed at the QB site leaves and is replaced by a

+ new plastoquinone molecule. The electron-deficient primary donor P680 is subsequently reduced via a four-step oxidation (S-state transitions) of water to molecular oxygen by the water-oxidizing complex (also known as the oxygen-evolving complex, OEC) that includes an Mn4-Ca cluster (reviewed in Bricker and Ghanotakis, 1998; Nelson and

Yocum, 2006; Kern et al., 2007). The OEC is oxidized by a redox-active and neutral tyrosine radical (TyrZ, Y161) located on the D1 polypeptide of the RC (Barry and

+ Babcock, 1987). Finally P680 is reduced and becomes available for another round of the light-driven charge separation.

PS II was crystallized as a dimer in which each monomer is characterized by a pseudo-twofold symmetry that equates the D1, CP43, and PsbI subunits to the D2, CP47, and PsbX subunits (Fig. 1.11). The RC polypeptides, D1 and D2, form the core of the complex, each consisting of five TM helices that structurally resemble the L and M chains of the bacterial reaction center. Flanking the RC core lie the proximal antenna subunits CP43 (PsbC) and CP47 (PsbB) comprising six transmembrane helices each. The six transmembrane spans of CP43 and CP47 are arranged in a manner similar to the six

N-terminal TM spans of the PsaA and PsaB subunits of PS I and have similar light- harvesting functions. Surrounding the RC core and proximal antenna are several low molecular weight membrane polypeptides including PsbE and PsbF that form the  and  subunits of cytochrome b559 (Cyt b559). Later crystallographic studies have also demonstrated the presence of lipids within the PS II complex which are concentrated in the region between the RC and proximal antenna (Loll et al., 2005; Guskov et al., 2009).

Owing to the composition and location of the lipid molecules, they have been implicated

39 in providing structural flexibility to the complex and aiding in the turnover of protein subunits (D1) during photooxidative stress.

Fig. 1.11 View of the PS II complex (Gokhale and Sayre, 2009). Helices are represented as cylinders with D1 in pink; D2 in violet; CP43 in orange; CP47 in firebrick red; Cyt b559 in grey, psbH in purple; psbI, psbJ, and psbK (barely visible) in cyan; psbL, psbM, psbN, and psbO in lime green; psbT, psbU, and psbV in yellow orange and psbX and psbZ are scarcely discernible in the backdrop in smudge green. The RC Chls are shown in light green, antenna Chls in forest green, pheophytins in blue, quinones in purple, β- carotenes in orange, non-heme iron is shown in red and the hemes in magenta. The oxygen atoms of the oxygen evolving complex are shown as spheres of light red, the manganese ions in purple, and the calcium ion in lime green.

The PS II RC core binds six chlorophylls (PD1, PD2, ChlD1, ChlD2, ChlzD1, and ChlzD2), two pheophytins (PheoD1 and PheoD2), two plastoquinones (QA and QB), two carotenoids

(CarD1 and CarD2) and a non-heme iron (Fig. 1.12). The cofactors are arranged in parallel

C2 symmetry-related branches with nearly identical cofactor orientations and center-to- center distances between cofactors. Interestingly, only one of these branches (D1 side) is active in electron transfer. It is hypothesized that local asymmetries in protein–cofactor interactions account for the unidirectionality of electron transfer in PS II.

Fig. 1.12 The organization of electron transfer cofactors in cyanobacterial PS II reaction center crystals (Loll et al., 2005, Nature 438: 1040 – 1044).

1.3.2 D1 and D2 RC polypeptides

Originally, many polypeptides of the PS II reaction center were identified through comparative analyses of profiles from wild-type and PS II mutant thylakoid membranes isolated from Chlamydomonas (Chua and Bennoun, 1975). The D1 and D2 polypeptides were first resolved using a continuous urea gradient which separated the bands migrating at ~30 kD into two distinct species (Satoh et al., 1983). The D1 polypeptide (diffuse band

1), as it came to be known, was proven to contain the QB herbicide-binding site through radiolabeling studies and shown as being rapidly synthesized and turned over in illuminated chloroplasts (Pfister et al., 1981; Grebanier et al., 1978; Satoh et al., 1983).

Subsequently, it was shown that the 32-kDa D1 RC protein was encoded by the chloroplast-encoded psbA gene (Zurawski et al., 1982). The gene encoding the D2 protein, psbD, was identified when an open reading frame was discovered in the chloroplast genome that was highly similar to the psbA gene of Chlamydomonas

(Rochaix et al., 1984). The D1 and D2 polypeptides showed 27% primary sequence identity and similar hydropathy plot profiles. Many of the conserved residues were found clustered near the cofactor-binding sites suggesting that they shared similar functions.

When the D1 and D2 amino acid sequences were revealed, their relatedness to the sequences of the L and M subunits of the Rhodobacter sphaeroides bacterial reaction center was quickly recognized (Hearst and Sauer, 1984). This observation became especially valuable when the type II bacterial RC (BRC) from Rhodopseudomonas viridis was crystallized and its structure determined at 3 Å and later 1.6-Å resolution

(Deisenhofer et al., 1985; Michel and Deisenhofer, 1988). The quinone-type BRC has

44 four subunits; H (heavy), M (medium), L (light), and cytochrome c. The L and M subunits each have five transmembrane helices and form the core of the complex. They coordinate four bacteriochlorophylls (BChlSP, BChlL, and BChlM), two bacteriopheophytins (BPheoL and M), two quinones, one non-heme iron, and one carotenoid. The H subunit has one TM helix and a globular domain associated with the

L/M heterodimer on the cytoplasmic side. The cytochrome subunit covalently binds four heme groups and is attached to the L/M complex on the periplasmic side. Upon photoexcitation, primary charge separation is initiated by the primary electron donor, a special pair of BChls (BChlSP). Electrons travel up the active (L) branch through intermediate cofactors finally reducing the first stable electron acceptor, a quinone. The oxidized BChlSP is re-reduced by cytochrome c. The sequence similarity between the D1 and D2 subunits and the L and M polypeptides is most conserved in the environment surrounding the reaction center chlorophylls and the non-heme iron. This information in conjunction with hydropathy plots was used to predict the topology of the D1 and D2 polypeptides in PS II (Trebst, 1986, 1987). The presence of five TM units per chain was confirmed by mapping D1 and D2 proteolytic fragments identified by antibodies generated against synthetic peptides (Sayre et al., 1986; Trebst, 1986). The isolation of a

PS II RC from spinach established that the D1 and D2 proteins bound the chlorins and quinones of PS II (Nanba and Satoh, 1987). Before the architecture of PS II was known, a number of homology-based structural models for D1 and D2 had been proposed based on similarities to the L and M subunits of the BRC (Svensson et al., 1990, 1996; Ruffle et al., 1992; Xiong et al., 1996). When the cyanobacterial PS II structure was solved, it was

45 unambiguously confirmed that D1 and D2 formed a heterodimer each with five TM spans and coordinated or bound several ET cofactors arranged in two parallel branches across the membrane. Unlike the BRC however, PS II possesses a tetra-Mn cluster capable of catalyzing the oxidation of water, yielding molecular oxygen. In addition, symmetry- related and redox-active tyrosine residues have been identified on the D1 (TyrZ) and D2

(TyrD) polypeptides. Other conspicuous differences between the BRC and PS II RC polypeptides are the residues that ligate or are located nearest to the Mg2+ atom of the monomeric chlorophylls (ChlD1 and ChlD2 in the PS II RC), being histidine residues in the

BRC and D1-T179 and D2-I178 in PS II. Also, D1 and D2 bind two additional chlorophylls (ChlZD1 and ChlZD2) located on their peripheries, which are coordinated by

D1-H118 and D2-H117 (Hutchison et al., 1995) (Fig. 1.13).

1.3.3 Primary electron donor, P680

PS II is unique in its capability of catalyzing the oxidation of water to molecular oxygen. This process is driven by the high redox potential (~+1.2 V) of the primary electron donor of PS II, also known as P680. (The name is based on the maximal Qy

+ absorption peak of the primary donor.) The P680/P680 midpoint potential is ~400 mV more positive than the midpoint potential of free chlorophyll a in solution (0.8 V for Chl a in dimethylformamide) and ~800 mV more oxidizing than the primary donor of the purple bacterial RC (Allen and Williams, 1998; Kato et al., 2009). In all other RCs the midpoint potential of the primary donor is more negative than free Chl a in dimethylformamide. Interestingly, the six chlorins located in the core of the reaction

46 center complex are nearly isoenergetic and are spectrally indistinguishable from each other (Ishikita and Knapp, 2005; Ishikita et al., 2005). These up-shifted values for redox potentials can by and large be attributed to the multiple cofactor–protein and cofactor– cofactor interactions involving the core Chls. It had been previously proposed that P680 is likely a multimer composed of a set of weakly coupled pigments including PD1, PD2,

ChlD1, ChlD2, PheoD1, and PheoD2 (Durrant et al., 1995). In spite of the obvious homology between the bacterial reaction center (BRC) and PS II RC, this model for the primary donor in PS II differs substantially from that of the BRC in which it has been shown that the primary donor is a BChl a special pair. By and large the organization of PD1 and PD2 is comparable to BChlSP of the BRC, however the relative orientation and center-to-center distance between their head groups differs slightly from their bacterial counterparts, the distance being longer (8.2 Å in 15SL and 7.6 Å in 2AXT) in PS II (Ferreira et al., 2004;

Loll et al., 2005). This difference may account for the weaker coupling between PD1 and

PD2 and the monomeric properties exhibited by P680 (Barter et al., 2003). The true identity of P680 is still unknown; however several lines of evidence suggest that P680 is the monomeric ―accessory‖ ChlD1. The lack of any significant red-shift in the Qy absorption band expected for an energetically coupled Chl dimer, and electron paramagnetic resonance (EPR) spectroscopic evidence for the localization a Chl triplet oriented 30° relative to the membrane plane normal (not consistent with the orientation of the ChlSP), suggest that the active branch monomeric Chl, ChlD1, is the probable primary electron donor in PS II (van Meighem et al., 1991; van Gorkom and Schelvis, 1993).

More recently, it has been shown that ChlD1 and not the ChlSP, has the lowest site energy and hence the excitation energy is more localized on this RC Chl, particularly at liquid helium temperatures (Raszewski et al., 2005; Schlodder et al., 2008a). This makes

* * ChlD1 , rather than ChlSP , the plausible primary electron donor and the site of initiation of primary charge separation in PS II. At ambient temperatures, however, the excited state is more delocalized over the entire multimeric set of RC chlorins that include the

ChlSP, ChlD1/D2, and PheoD1/D2, but primary charge separation is still initiated from ChlD1

(Durrant et al., 1995; Dekker and van Grondelle, 2000; Barter et al., 2003). Flash induced

+ - 3 1 (P680 QA —P680QA) and ( P680- P680) absorbance difference spectra of the D1-H198Q and

D1-T179H Synechocystis mutants with respect to the WT, have indicated that the RC

3 + - triplet that arises from P680 PheoD1 recombination is localized on ChlD1 whereas the oxidized primary donor cation is localized on the PD1 Chl of the PS II ChlSP (Schlodder et al., 2008b). In addition, recent reports have found that the reduction of PheoD1 occurs prior to the oxidation of PD1/PD2 of the ChlSP, even at ambient temperatures, implying that the cation of the primary radical pair is localized at a site other than the ChlSP, most likely

ChlD1 (Groot et al., 2005 and Holzwarth et al., 2006).

Site-directed mutations of the D1-H198 and D2-H197 residues that serve as ligands for the PS II RC ChlSP have also been very useful in deciphering the organization of P680.

The substitution of the PD1 ligand with glutamine has been shown to induce a blue shift in the Chl Soret band and Qy transitions and alter the kinetics of electron flow between P680 and QA as well as between TyrZ and P680 suggesting a modification of the redox potential of P680 (Diner et al., 2001). The kinetic and spectral effects of mutating the Mg axial

+ ligand to PD2 were much less severe, suggesting that P680 cation is localized primarily on

PD1 of the ChlSP. The substitution of the histidine ligands of PD1 and PD2 to non- coordinating residues like alanine and leucine were surprisingly unremarkable and and led to the formation of reaction centers that functionally resembled those of the wild type

(Diner et al., 2001). It was reasoned that a water molecule provided the Mg2+ atom the axial ligand necessary for PD1 coordination in the mutant reaction centers. However, in the case of substitutions with larger non-ligating residues like Leu and Tyr, the mutant reaction centers failed to assemble possibly due to the displacement of the proposed coordinating water molecule (Vermaas et al., 1988a; Diner et al., 2001). A similar type of coordination was seen by water in the reaction centers of L-H173G and M-H200G mutants of R. sphaeroides. Substitution of the PD1 and PD2 axial ligands with Leu did not, however, corroborate results from analogous replacements in the bacterial reaction center, where the reaction center assembled but with the incorporation of BPheo in the place of the BChls of the special pair.

A greater degree of asymmetry exists between the axial ligands for the RC Chls in PS

II when compared to those in the BRC. More specifically, in the BRC, all the axial ligands to the RC BChls are histidine residues. On the other hand in the PS II RC, while the axial ligands of the PD1, PD2, ChlZD1 and ChlZD2 RC Chls are histidines, the active and inactive branch monomeric Chls, ChlD1 and ChlD2, lack protein ligands (Fig. 1.13).

Instead, the amino acid residues D1-T179 and D2-I178 are found to be in closest proximity to ChlD1 and ChlD2 respectively (Ferreira et al., 2004). In Chapter 3 of this thesis, we describe results of modifying the local protein environment of ChlD1 to

49 elucidate its function in charge separation as well as to study the effects of the proximal amino acid residues on its redox properties. Our results show that the nature of the residue in position 179 of the D1 RC-polypeptide (closest to ChlD1) is involved in mediating excitation energy transfer to the RC and is paramount in determining the redox properties of the primary electron donor. The latter plays a critical role in balancing forward ET with charge recombination.

Fig. 1.13 PS II RC Chls and the surrounding protein environment (Gokhale and Sayre,

2009).

1.3.4 Primary electron acceptor, PheoD1

Originally, the bound plastoquinone in PS II, QA, was thought as being the primary electron acceptor in quinone-type RCs. However, in the late 1970s it was discovered that the principal photochemical event is marked by ET from the excited primary donor to

PheoD1, located on the active branch (Klimov et al., 1977; Klimov and Kravnoski, 1981;

Klimov, 2003). Recently the redox potential of PheoD1 was determined using thin-layer cell spectroelectrochemistry as being ~-505  6 mV (Kato et al., 2009), which was ~100 mV more positive than what was previously thought (Klimov, 1979). In isolated bacterial reaction center preparations, primary charge separation occurs in 2.8 ps at room temperature and 0.7–1.2 ps at 10 K (Woodbury et al., 1985; Martin et al., 1986). Primary charge separation in PS II RCs using sub-picosecond time resolution, has been shown to occur with a time constant of 3.0  0.6 ps at 277 K (Wasielewski et al., 1989). Formation

+ - of the charge-separated state (P680 PheoD1 ) was monitored by transient absorption spectroscopy between the wavelengths of 450 and 840 nm following excitation at 610

+ nm. The appearance of an absorption band at 820 nm indicated the formation of P680 and was accompanied by absorption changes at 505 and 540 nm due to the formation of the

PheoD1 anion. The value of the time constant for the appearance of the charge-separated state was comparable to the time constant for the decay of the lowest singlet excited state of P680. However, a different rate constant of 21 ps for primary charge separation has also been interpreted from similar ultrafast experiments carried out by Durrant et al. (1992,

1993). This calculation was based on the observed rate of bleaching of the pheophytin ground state at 545 nm, the appearance of the pheophytin anion at 460 nm, and the loss of

52 stimulated emission from Chl proposed to be caused by the formation of the oxidized primary donor. The kinetics of charge separation when derived based on absorption changes in the 545 nm region (PheoD1 QX band), is multiphasic in nature and can be exponentially fit to yield three lifetime components of 3 ps, 20 ps, and between 50ps and

100 ps (Greenfield et al., 1997). The slower (50 ps) lifetime components have been assigned to charge separation that is limited by energy transfer processes with the RC

(―equilibration‖).

Data from femtosecond visible-pump–mid-infrared probe spectroscopy on PS II RCs in the region of the chlorophyll ester and keto modes, between 1,775 and 1,585 cm-1 showed that showed that while the reduction of pheophytin occurs on a 0.6- to 0.8 ps time

+ scale, the PD1 cation, is formed only after 6 ps (Groot et al., 2005). The authors concluded that PheoD1 is the primary electron acceptor in PS II and that the first charge separation step occurs between the ‗‗accessory chlorophyll‘‘ ChlD1 and the active PheoD1.

More recently, femtosecond transient absorption studies probing the spectral region of the

Pheo Qx band (540–550 nm) in isolated PS II RCs and intact PS II cores, showed that energy equilibration between antenna and RC occurred in ~1.5 ps, while the apparent lifetime of primary charge separation between ChlD1 and PheoD1 was ~5.5 ps in intact PS

II cores (Holzwarth et al., 2006). The 35-ps component represented the apparent lifetime

+ - of formation of a secondary radical pair (PD1 PheoD1 ). In isolated RCs, the apparent lifetimes of primary and secondary charge separation were found to be 3 and 11 ps, respectively. The data showed that pheophytin is the primary electron acceptor in PS II and is reduced in first ET step, and that the rate constants of electron transfer in the RC

53 are identical for PS II cores and for isolated RCs. The accessory Chl, ChlD1 was proposed as being the primary electron donor.

Biochemical analyses as well as evidence from crystal structures have revealed that

PS II has two pheophytins per reaction center which can be distinguished on the basis of their optical properties. The active-branch pheophytin has an absorption peak centered at

676 nm whereas the inactive-branch pheophytin contributes to the peak at 680 nm

(Breton, 1990; Tang et al., 1990; van Kan et al., 1990; van Gorkom and Shelvis, 1993).

The two pheophytins are related pseudo-symmetrically but there is a clear difference in the number of hydrogen bonding residues that surround them. For instance, in the eukaryotic PS II RC, a strong hydrogen bonding interaction exists between the ring V carbonyl group of the active branch PheoD1 and D1-E130 (D1-Q130 in cyanobacteria)

(Moënne-Loccoz et al., 1989; Dorlet et al., 2001). Two additional putative H-bonds may be provided by D1-Y126 and D1-Y147, one to the PheoD1 head group and the other to the phytyl tail which are lacking in the environment surrounding PheoD2. The analogous inactive branch residues are D2-Q129, D2-F125 and D2-F146, whose side-chains are in comparison less capable of hydrogen bonding interactions (Fig. 1.14).

High field EPR studies of the radical anion of PheoD1 from the Chlamydomonas D1-

E130 mutants (Q, H, and L) showed a strong effect of the hydrogen bond strength on the g tensor of the Pheo radical (Dorlet et al., 2001). The D1-E130Q and L mutants showed upward shifts in gx value as compared to wild type, consistent with a weakening or loss of the hydrogen bonding interaction. Subsequently, Chl fluorescence decay kinetic data obtained for the D1-E130L Chlamydomonas and D1-Q130L cyanobacterial mutants

54 showed that the loss of the hydrogen bond interaction to PheoD1 resulted in a longer

- lifetime for S2QA charge recombination (Cuni et al., 2004; Rappaport et al., 2005; Cser and Vass, 2007). Additionally, thermoluminescence measurements on the same mutants

+ - + - revealed that the energetic gap between the primary radical pair P680 PheoD1 and P680 QA increased as a result of a leucine substitution and the loss of a hydrogen bond donor to

PheoD1 (Rappaport et al., 2005; Cser and Vass, 2007). It was inferred that mutagenesis of

D1-130 to amino acids that weakened the hydrogen bonding interaction to PheoD1 shifted

- the midpoint potential of the PheoD1/PheoD1 couple to more negative values, reducing the probability of forward ET transfer from PheoD1 to QA. Hence, the D1-E130L PS II mutation effectively results in an impairment of forward ET and reduction in primary charge separation yield. In nature, the cyanobacterial D1 protein exists in two conformations: a low light version in which a glutamine residue is present in position 130 of the D1 protein and a high light version containing glutamate (Clarke et al., 1993).

Switching from the low to the high light form causes an increase in the strength of the hydrogen bonding interaction between D1-130 and the PheoD1 head group, leading to a

+ - + - decrease in the free energy gap between P680 PheoD1 and P680 QA and a consequent increase in the quantum efficiency of photosynthesis. The switch from the low to high light form of D1 also decreases the yield of the indirect radiative charge recombination

3 * 3 1 pathway in which P680 is formed, thereby reducing P680-mediated O2 formation and photooxidative damage (Krieger-Liszkay et al., 2008; Vass and Cser, 2009).

As mentioned previously, PheoD2 is related to PheoD1 by C2-symmetry but does not take part in primary ET in PS II. The role of PheoD2 and its protein environment is not as

55 well understood as the role and protein environment of PheoD1. There is some evidence to show that PheoD2 may be excitonically coupled to the central multimeric RC chlorins and hence be involved in excitation energy equilibration within the RC complex (Durrant et al., 1995; Xiong et al., 2004). The substitution of PheoD2 with a chlorophyll (Chl) molecule has been shown to result in substantial impairment of PS II due to the possible redistribution of the excited-state equilibrium among the core pigments of the PS II RC

(Xiong et al., 2004). The inactive branch residue analogous to D1-E130 in chloroplastic

PS II is D2-Q129. D2-Q129 has also been suggested as being a possible hydrogen bond donor to the head group of PheoD2 (Xiong et al., 1996; Kern and Renger, 2007).

However, analysis of the most recent PS II crystal structure from the cyanobacterium

Thermosynechococcus elongatus at 2.9 Å resolution reveals that the side chain of D2-

Q129 is 3.9 Å away from the PheoD2 head group and therefore may not be in close enough proximity to serve as a viable hydrogen bond donor (Fig. 1.14). In Chapter 4, we describe the role of the highly conserved acceptor side residue D2-Q129. We use a mutagenesis approach in which D2-Q129 was replaced by a non-conservative hydrophobic leucine residue (D2-Q129L) or a conservative histidine residue (D2-Q129H) in Chlamydomonas reinhardtii. Our results demonstrate that the amino acid residue in position 129 of the D2 protein has a major influence on the redox potential of QB. This result was unanticipated given that the analogous active branch residue, D1-E130, influences the redox properties of PheoD1. The disparate effects of the analogous D1-

E130L and D2-Q129L Chlamydomonas mutants are detailed in Chapter 4, and a model

56 for the structural impacts of the D2-Q129 mutations on the QB site and on the redox potential of QB is proposed.

Fig. 1.14 Protein environments of PheoD1 and PheoD2 in PS II. Relevant distances between atoms are shown as dashed lines. Figure was created using PyMOL (Delano

Scientific) using 3BZ1 PDB coordinates.

1.3.5 Secondary electron acceptors, QA and QB

During light-driven charge separation a plastoquinone molecule in the QA site, typically a single electron acceptor, accepts an electron from PheoD1. Due to the higher midpoint potential of QA than PheoD1, the charge-separated state is stabilized, reducing the probability of a back reaction (Fig. 1.15). Under high light conditions, however, QA can accept two electrons and become doubly reduced, prompting the turnover of the D1

RC protein (Keren et al., 1995). Electron flow from QA to QB is considered the rate- limiting step of PS II ET and occurs on the microsecond time scale.

Fig. 1.15 Energetics for electron transfer components in PS II based on the redox potentials (Kato et al., 2009, Proc. Natl. Acad. Sci. U. S. A. 106: 17365 – 17370).

Although PS II and the BRC possess different quinone molecules, their binding pockets are structurally similar. The architecture of the QA site is well elucidated by the crystal structures available for the cyanobacterial PS II and consists mainly of hydrophobic residues belonging to the D2 polypeptide including D2-I213, H214, T217,

Y244, M246, A249, N250, W253, A260, F261 and L267 (Ferreira et al., 2004; Loll et al.,

2005; Kern and Renger, 2007) (Fig. 1.16). In addition, the keto-oxygens of the QA head group are hydrogen bonded to the side chain of D2-H214 and the main chain amide nitrogen of D2-F261 (Loll et al., 2005). The terminal end of the isoprenoid chain of QA lies between the phenyl rings D1-F52 and PsbT-F10 (Guskov et al., 2009). The QA site residues are that conserved include D2-T217, D2-F252, D2-W253, and D2-H214 which correspond to M-T220, M-F249, M-W250, and M-H217 of the BRC.

The QB binding site in PS II is formed exclusively by D1 protein residues and features a slightly more open pocket than that for QA binding. The QB site in PS II is less conserved in comparison with the BRC than the QA site. Three structural elements define the QB site namely the C-terminal part of TMH d, spanning residues D1-G207-V219; the cytosolic (stromal) surface helix (D1-de helix) and the connecting D1-de loop region; and lastly, the N-terminal part of TMH e, spanning residues D1-R269-F274 (Kern and

Renger, 2007). The QB keto-oxygens are hydrogen bonded to the side chains of D1-

H215, S264 which correspond to the L-H190 and L-S223 in the BRC. An additional hydrogen bond to the backbone amide of D1-F265 is present (Ferreira et al., 2004; Loll et al., 2005). Like the QA binding pocket, the QB pocket also houses mainly hydrophobic

61 residues which include D1-M214, H215, L218, V219, Y246, I248, A251, H252, F255,

S264, F265 and L271 (Kern and Renger, 2007; Takahashi et al., 2010) (Fig. 1.16).

In addition to plastoquinone, the PS II QB binding pocket can bind a variety of herbicides namely, atrazine, 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), bromacil, and metribuzin. Hence, the QB site been studied extensively in relation to herbicide binding and resistance. Spontaneous or chemically induced herbicide resistance mutations have been shown to be clustered in the QB binding cavity (residues 211–275 of the D1 protein) and have variable impacts on the rate of QA to QB electron transfer

(Erickson et al., 1989; reviewed in Diner et al., 1991; Crofts et al., 1993). For instance, the D1-S264A mutant exhibits increased tolerance toward metribuzin and atrazine but slower QA → QB ET than the wild type. Additionally, the D1-G256D mutation confers

- - atrazine resistance but also slows QA → QB ET (Crofts et al., 1993). Other mutations of the QB-binding pocket such as D1-A251V produce atrazine-resistant phenotypes with

- slower QA → QB ET rates, whereas D1-V219I, D1-F255Y, and D1-L275F herbicide

- - - tolerant mutants show only modest changes in QA → QB and QA → QB ET rates

(Johanningmeier et al., 1987; Erickson et al., 1989; Govindjee et al., 1991; Strasser et al.,

1992).

Several studies involving the site-specific mutagenesis of the QB binding D1 protein

Chlamydomonas and Synechocystis have also led to the identification of D1 residues involved in herbicide binding (Przibilla et al., 1991; Ohad and Hirschberg 1990, 1992;

Kless et al., 1994; Mäenpää et al., 1995; Xiong et al., 1997; Lardans et al., 1998). For example, site specific mutagenesis of the C. reinhardtii D1 protein at positions 259, 264

62 and 266 to generate the double (D1-N266T/D1-I259S) and triple (D1-N266T/D1-

S264A/D1-I259S) was found to provide metribuzin-resistance (Przibilla et al., 1991).

Similarly, replacements of the QB-binding site residue, D1-A251, with 12 different amino acids resulted in some herbicide resistant mutants. Five substitutions (Arg, Asp, Gln, Glu, and His) led to the impairment of photosynthesis. Whereas the D1-A251R mutant was able to synthesize a less stable but full-length 32-kD D1 protein, the other four mutants produced truncated (24–25 kD) and unstable versions of the D1 protein (Lardans et al.,

1998).

- - PS II mutants of the QB binding pocket often have altered QA QB ↔ QAQB equilibrium arising from a change in the redox potential of QB, as monitored by Chl fluorescence decay and/or thermoluminescence measurements (Ohad and Hirschberg,

1992; Etienne and Kirilovsky, 1993; Kless et al., 1994; Nixon et al., 1995; Constant et al.,

1996; Xiong et al., 1998; Lardans et al., 1998; Minagawa et al., 1998; Sane et al., 2002;

Rose et al., 2008). Interestingly, the mutations described in the aforementioned studies are in the D1 protein of the PS II reaction center complex. To our knowledge, only one study exists thus far in which multiple mutations of the D2-de loop of the D2 protein alter

- - the redox potential of QB and hence the QA QB ↔ QAQB equilibrium (Kless et al., 1993).

In Chapter 4, we discuss the effects of a point mutation of the D2 polypeptide, at residue

D2-Q129, which leads to the modification of QB redox potential. Our data show that this unique mutation of the D2 protein decreases in the free energy gap for recombination of

- QB with S2 but does not impact the redox potential of QA or of the donor side cofactors.

- - Additionally, we show that D2-Q129 mutagenesis causes a shift in QA QB ↔ QAQB

- equilibrium toward QA , and increases susceptibility to photoinhibitory light due to

- 3 increased S2QB charge recombination and P680-mediated oxidative damage.

The most recent and best resolved crystal structure of cyanobacterial PS II reveals the presence of a third plastoquinone molecule (QC) located near the QB site (Guskov et al.,

2009). However due to its limited electron density, it could be modeled with only five isoprenoid units. It is seen to form van der Waals contacts with CarD2, the isoprenoid chain of QB and the phytol chain of ChlD2, but does not interact with protein. It is situated

~17 Å away from QB and ~15 Å from the heme of cyt b-559 and has been implicated in quinone exchange as well as in the regulation of PS II function in cooperation with cyt- b559. However, its true function is still unknown.

- The presence of a non-heme iron in the proximity of QA was predicted early on by

- EPR measurements detecting the QA /Fe(II) signal and was later confirmed by crystallographic data to be 9 Å apart (Fig. 1.11) (Nugent et al., 1981; Ferreira et al., 2004;

Loll et al., 2005). The non-heme iron mediates ET between QA and QB and is coordinated by four histidine residues D1-H215, D1-H272, D2-H214, and D2-H268, two each from the D1 and D2 proteins (Fig. 1.16). In contrast to the BRC, where a glutamate residue provides a fifth Fe ligand, bicarbonate serves the same function in PS II (Ferreira et al.,

2004; Loll et al. 2005; Guskov et al., 2009). In addition, it has been suggested that at physiological pH, the bicarbonate ion forms a hydrogen bonding network with the neighboring D1 and D2 amino acid residues helping to stabilize the complex and aid in

ET (reviewed in van Rensen et al., 1999). It has also been shown to aid in the protonation

- of QB (Xu et al., 1991). The displacement of bicarbonate by formate decreases the

64 kinetics of QB reduction by 100-fold. The formate/bicarbonate effect has been studied by monitoring EPR spectra of the QA-Fe-QB complex (Bowden et al., 1991). Prior to the localization of the bicarbonate-binding site by crystallography, attempts were made to localize it by site-directed mutagenesis. Positively charged residues located near the non- heme Fe, such as D1-R269, were targeted for mutagenesis. A non-conservative glycine mutation of D1-R269 was engineered into Chlamydomonas (Hutchison et al., 1996b).

The D1-R269G mutant was unable to evolve oxygen, had an increased susceptibility to photoinhibition and lacked the manganese associated with the water-splitting apparatus.

Additionally, Chl a fluorescence decay kinetics revealed a blockage in electron transfer from QA to QB that was thought to be consistent with loss of bicarbonate binding. In

- dark-grown cells, the TyrD and QA /Fe(II) EPR signals were only 30% that of wild-type cells indicating reduced levels of functional PS II complexes in the mutant. However, the

- amplitude of the formate-enhanced QA /Fe(II) EPR signal was consistent with the predicted levels of functional PS II complexes in the membrane suggesting that the mutant was capable of binding bicarbonate and that the negative effects on forward electron transfer yield and assembly of the water-splitting complex, was due to structural changes caused by the stromal side mutation.

Fig. 1.16 View of the non-heme iron and the QA and QB binding pockets (Ferreira et al.,

2004, Science 303: 1831 – 1838).

1.3.6 Oxygen evolving complex (OEC)

The catalyst for water oxidation, the OEC, is the charge-accumulating tetra manganese complex which is bound to the D1 and CP43 proteins. The complex is stabilized by chloride and calcium ions and the three extrinsic small subunits (PsbO,

PsbQ and PsbP in green algae and higher plants and PsbO, PsbU and PsbV in cyanobacteria) (Mayfield et al., 1987a,b, 1989; Bricker and Ghanotakis, 1996; Nelson and Yocum, 2006; Roose et al., 2007). The driving force for water oxidation comes from

+ P680 which oxidizes TyrZ, which in turn oxidizes the tetra-Mn complex. The oxidation of two molecules of water requires four quantum events which drive the OEC through five redox transitions or S-states (S0 → S4) beginning with the dark-stable S1 state (Kok et al.,

1970). Moreover, calcium and chloride have been shown to be required for the photoactivation or the light dependent assembly of the tetra-Mn complex (Homann,

1988).

The shape of the electron density attributed to the Mn4Ca cluster in the 3.0 Å (Loll et al., 2005) and the 2.9 Å PS II structure (Guskov et al., 2009) are best approximated by four Mn ions arranged in the form of a hook. The Mn ions are numbered Mn1 to Mn4, with the lowest number attributed to the highest electron density (Fig. 1.17). This geometry is reminiscent of the Y-shaped arrangements proposed based on the 3.8 Å

(Zouni et al., 2001) and 3.2 Å (Biesiadka et al., 2004) structures but are markedly different from the 3.5 Å structure (Ferreira et al., 2004) that proposed a cubane-like

Mn3CaO4 cluster linked to a fourth Mn by a mono-µ-oxo bridge. In the 2.9 Å crystal structure, Mn1 and Mn2 (Mn1-Mn2) and Mn2 and Mn3 (Mn2-Mn3) are seen to be

67 connected by di-µ-oxo bridges and are 2.7Å apart; while Mn1 and Mn3 (Mn1–Mn3) and

Mn3 and Mn4 (Mn3–Mn4) are placed 3.3 Å apart suggesting that they are probably connected by mono-µ-oxo bridges (Loll et al., 2005; Guskov et al., 2009). The hydroxyl group of YZ is 5 Å apart from the Ca and at distances of 7.5-8.3 Å to the four manganese.

2+ Interestingly, the Ca is appropriately positioned between Mn4 and YZ, implying an important role in proton coupled electron transfer between the Mn4Ca cluster and YZ and water oxidation. The depletion of calcium from PS II has been shown to prevent oxygen evolution and results in the loss of the normal g = 2 multiline EPR signal and its replacement by a unique dark stable multiline spectrum (Dorlet et al., 1999). Recent EPR studies have indicated that the blockage caused by calcium depletion in forming the S3 state may be overcome at low pH, suggesting that calcium depletion might disrupt the

• proton-conducting pathway to YZ upon its reduction by the OEC (Styring et al., 2003).

The location of chloride in PS II was revealed by the 2.9 Å crystal structure and is found bound by water(s) to the Mn4Ca cluster (Fig. 1.17). It is located 6.5 Å from the

Mn4Ca cluster and coordinated by D2-Lys317NZ, D1-Asn181ND and D1-Glu333N (Fig.

1.17) (Guskov et al., 2009). On the basis of oxygen-evolution and EPR measurements in the presence of other anions (bromide, iodide, nitrate, fluoride) in comparison with chloride, the role proposed for chloride in water-splitting was to facilitate proton transfer from the manganese cluster to the lumen, along with other charge amino acids (Olsen et al., 2003). Unlike calcium, however, chloride may be replaced by other anions such as bromide and iodide, though oxygen evolution becomes less effective (Homann, 1988;

Olsen et al., 2003). A modified S2 state can be formed in chloride-depleted PS II

68 complexes, but it does not advance without re-addition of chloride (Sandusky and

Yocum, 1986; Homann, 1988).

The coordination sphere of the Mn4Ca cluster is provided by the amino acid residues from polypeptide D1 and CP43. The PsbO, PsbV and PsbU extrinsic proteins that are involved in stabilizing the complex, do not serve as direct ligands to the metals ions.

Based on the 2.9 Å PS II crystal structure it was proposed that five carboxylate residues

(D1-E189, D1-D342, D1-E333, CP43-E354, D1-A344) possess the ability to act as bidentate ligands bridging different cations of the OEC, enhancing their stability and facilitating the S-state transitions (Fig. 1.17) (Guskov et al., 2009). D1-E189 and D1-

A344 in D1 have also been identified as possible ligands for Ca2+.

The direct oxidant of the OEC Mn cluster is tyrosine-161 of the D1 subunit, YZ, which was previously suggested to act as a redox-coupled base (Babcock et al., 1989;

+ Britt, 1996). According to this model, it was suggested that the oxidation of TyrZ by P680 generates a neutral tyrosine radical alongside the exchange of a proton between TyrZ and

D1-H190. The D1-H190 residue functions as a weak base which shuttles a proton from

TyrZ to the thylakoid lumen (Tang et al., 1994; Britt, 1996; Babcock et al., 1986). This stepwise reduction and protonation of the oxidized TyrZ was thought to generate higher

S- (oxidation) states of the tetra-Mn complex leading to the oxidation of water, presumably located near the calcium-binding site and Mn4. Support for this model came from EPR studies that demonstrated that oxidized TyrZ was a neutral radical, and from mutagenesis of D1-H190 in Chlamydomonas and later cyanobacteria that demonstrated its critical role in water oxidation. Chlamydomonas D1-H190 mutants are unable to

69 evolve oxygen and cannot assemble a tetra-Mn cluster (Roffey et al., 1994). The D1-

H190 mutants were also unable to generate the AT thermoluminescence band which is attributed to oxidation of TyrZ (Kramer et al., 1994). Structural homology models had predicted that D1-H190 was located near TyrZ, however D1-H190 mutations had no

• effect on the TyrZ EPR hyperfine structure. These results were in contrast to those observed with the symmetry-related D2-H189 residue. Non-conservative mutations of

D2-H189 altered the EPR-detectable hyperfine structure of TyrD associated with the loss of the strong hydrogen bond (Vermaas et al., 1988b; Tommos et al., 1993; Tang et al.,

1996). These results indicated that the H-bond distance between D1-H190 and TyrZ is greater than the H-bond distance between D2-H189 and TyrD. This prediction was later supported by the crystal structure. Subsequently, it was shown that TyrZ oxidation and proton release was a concerted reaction in wild-type PS II complexes but not in D1-H190 mutants (Hays et al., 1999).

Contrary to the hydrogen-abstraction hypothesis for YZ and D1-H190, it was proposed that during the S-state transitions, the same proton remains near YZ, moving only a short distance between YZ and D1-H190 depending on the redox state YZ. Support for this ―proton-rocking‖ hypothesis has been obtained from optical spectroscopy and proton release measurements using Mn depleted PS II enriched fragments in which the

ox - rate of the recombination reaction of YZ with the reduced primary acceptor QA was observed as being modulated by a protonizable group with pKa of ~6. Further, the same group was found to modulate the extent of proton release, releasing 1 per center above pH 7 and 0 below pH 5 (Rappaport and Lavergne, 1997; reviewed in McEvoy and

Brudvig, 2006). This proton rocking mechanism is thought to allow TyrZ to maintain a favorable redox potential for oxidation of the tetra-Mn complex. The restrained pairing between the YZ phenol group and the imidazole of D1–H190 side-chain allows for the rapid loss of a proton from the phenol group of tyrosine to D1–H190 upon tyrosine oxidation, and fast re-binding of the same proton upon reduction of the tyrosine by the

OEC.

It has recently been suggested that the arginine residue CP43-R357, located in close proximity to the OEC, plays the role of the redox-coupled base similar to the role that was proposed for TyrZ in the hydrogen-abstraction mechanism (McEvoy and Brudvig

2004; Brudvig, 2008). During the later S-states, oxidation of YZ decreases the pKa of

CP43-R357 such that its guanidinium side chain undergoes deprotonation. This electrostatic effect on CP43-R357 is achieved by the positive charge maintained at the

YZ/D1-H190 pair as described in the proton-rocking hypothesis. Recently, a computational study showed that the pKa of CP43-R357 is dramatically lowered by oxidation of the OEC from S2 to S3 (Ishikita et al. 2006). Additionally, a series of acidic and basic amino acid residues (D1-Asp61, D1-Glu65, D2-Glu312, D2 Lys317 D1-Asp59,

D1-Arg64, PsbO-Arg152, and PsbO-Asp224) were seen as forming a proton transfer pathway leading away from the CP43-R357 residue into the lumenal protein surface providing more evidence for the function of CP43-R357 as a base catalyst in the OEC

(McEvoy and Brudvig 2004; Brudvig, 2008). The proximity of CP43-R357 to the Ca2+ binding site and to a probable proton channel and the lack of proton channel leading from

TyrZ to the lumen, favors the base-catalyst role for the CP43-R357 residue in proton extraction from water.

Mutagenesis of CP43-R357 to a serine impairs but does not block oxygen evolution, and is associated with an 11-fold larger Kok double miss parameter and accelerated decay of advanced S-states (Ananyev et al. 2005). Unlike wild-type thylakoids, however, oxygen evolution in CP43-R357S is insensitive to bicarbonate depletion. Based on these observations it was suggested that CP43-R357 binds bicarbonate, which is required for oxygen evolution. It should be noted, however, that the magnitude of the bicarbonate effect on oxygen evolution in wild-type membranes is less than the magnitude of the effect of the CP43-R357S mutation. Thus, the CP43-R357S mutation may mask the effect of bicarbonate depletion in this mutant. The CP43-R357S mutant phenotype also has implications for the CP43-R357 base catalyst, metallo-radical water oxidation model.

The non-conservative CP43-R357S mutant would not be expected to function as a base catalyst for water oxidation.

Recent polarized X-ray absorption fine structure (EXAFS) data obtained for single crystals of PS II isolated from T. elongatus, yield at least four different arrangements for

+ the Mn4Ca2 cluster (Yano et al. 2006) which were all different than those proposed from

X-ray diffraction studies (Ferreira, et al., 2004; Loll et al., 2005). Radiation damage that can alter the oxidation states of the Mn ions during the collection of X-ray diffraction data has been suggested as being the root for inconsistencies between the different models (Yano et al. 2005). Although the exact geometry and protein environment of the

Mn4Ca cluster is still not known precisely, the proposed models provide a basis for

72 modeling the water oxidation process and dioxygen formation (reviewed in Barber,

2008). The knowledge of the location of one Mn ion (Mn4) adjacent to the Ca2+ and their relation to the side chains of key amino acids including D1-Y161 (TyrZ), D1-H190 and

CP43-R357, suggests that those two ions provide binding surfaces for the two substrate water molecules and water oxidation catalysis. One well-acknowledged mechanism suggests that the substrate water associated with Mn4 undergoes deprotonation during the

S-state cycle in a mechanism that is dependent on Mn4 being converted to a higher oxidation state possibly Mn(V) just prior to O–O bond formation (Messinger 2004;

McEvoy and Brudvig 2004, 2006). The electrophilic oxygen of the MnVO intermediate is then thought to undergo a nucleophilic attack by the oxygen of the second substrate water bound to Ca2+. An alternative mechanism suggests that the deprotonated water molecule on Mn4 forms an oxyl radical which attacks an oxygen atom linking Ca2+ with

Mn or the oxygen of a water molecule bound to the Ca2+ to form the O–O bond

(Siegbahn 2006).

Fig. 1.17 The Mn4Ca cluster of the oxygen evolving complex of PS II (Guskov et al.,

2009, Nat. Struct. Mol. Biol. 16: 334 – 342). 74

1.3.7 ChlZ cycle

Brudvig and coworkers (Thompson and Brudvig, 1988) first proposed that a

+ chlorophyll monomer known as ChlZ could reduce P680 with low quantum yield. The

ChlZ cation is re-reduced by Cyt b-559, which in turn is reduced by QB. In addition, a bound PS II carotenoid (CarD2) participates in the ChlZ electron transfer cycle around PS

II (Vrettos et al., 1999). The ChlZ cycle has been proposed to reduce photoinhibition and the light-dependent turnover of D1. Consistent with this idea, it has been shown that the

ChlZ cation is a potent quencher of PS II chlorophyll fluorescence (Schweitzer and

Brudvig, 1997; Schweitzer et al., 1998). The peripheral chlorophylls, ChlZD1 and ChlZD2, are coordinated by D1-H118 and D2-H117. Significantly, these residues and chromophores are not conserved in the BRC. Before the availability of the PS II crystal structure, it was demonstrated by site-directed mutagenesis that D1-H118 and D2-H117 indeed coordinated two additional Chls not present in the BRC (Hutchison and Sayre,

1995; Lince and Vermaas, 1998; Stewart et al., 1998). The location and identity of the

ChlZ cation is of particular interest because of its potential involvement in energy funneling and participation in the cyclic electron transfer pathway around PS II. In principle, both of the symmetry-related peripheral Chls are photooxidizable. However,

• EPR studies have indicated that only one ChlZ cation is accumulated per YD (Schweitzer et al., 1998). Therefore, it is likely that only one of the two peripheral accessory chlorophylls participates in the electron transfer cycle around PS II. Stewart et al. (1998) proposed that the chlorophyll coordinated by D1-H118 is the ChlZ species. This conclusion was based on the observation that the ChlZ resonance Raman spectrum was

75 altered in D1-H118 mutants but not in D2-H117 mutants. The PS II model structures

(Xiong et al., 1996) indicated, however, that the Chl coordinated by the D2-H117 residue was closer (25 Å) to QB and Cyt b-559 and that the D1-carotenoid was most closely associated with the chlorophyll special pair. Since QB indirectly reduces ChlZ, it is to be expected that the ChlZ peripheral chlorophyll would be nearer to QB than to QA.

Consistent with this speculation, Shigemori et al. (1998) demonstrated that the distance

• between YD and the ChlZ cation (29 Å) was consistent with the coordination of ChlZD2 by the D2-H117. Moreover, the efficient oxidation of Cyt b-559 by ChlZ suggested that ChlZ was in close proximity to the heme of Cyt b-559 (de Paula et al, 1985; Thompson and

Brudvig, 1988). Finally, Hutchison and Sayre (1995) and Wang et al. (2002) substituted the symmetrically located D1-H118 and D2-H117 with conservative amino acids (D1-

H118Q, D2-H117N, and D2-H117Q) in Chlamydomonas. They demonstrated that the

D2-H117 mutants had a reduced capacity to quench chlorophyll fluorescence (F695).

This was interpreted as being due to reduced ChlZ levels relative to wild-type and D1

H118Q mutant PS II complexes. Finally, the D2-H117 mutants also had reduced light- harvesting efficiency relative to wild-type and the D1-H118Q mutant (Johnston et al.,

2000; Ruffle et al., 2001). As a result, the D2-H117 mutants were less sensitive to photoinhibitory light treatment than wild-type and the D1-H118Q mutant. In summary, the accumulated structural and biophysical studies support the assignment of the ChlZD2 pigment as the chlorophyll monomer involved in the ChlZ cycle.

More recently, the oxidation pathway of ChlZ was studied at cryogenic temperatures by light-induced Fourier transform infrared (FTIR) difference spectroscopy (Kitajima and

Noguchi, 2006). The bleaching of the two Car molecules in the RC (CarD1 and CarD2) in

Mn-depleted PS II membranes at 250 K was monitored by observing the disappearance of light-induced FTIR signals of Car+ at 1465, 1440, and 1147 cm-1 under oxidative

+ -1 conditions. The ChlZ /ChlZ signal at 1713/1687 cm was observed even in Car-bleached

PS II under illumination at 80 K and had ~80% of the intensity of the control sample. The

+ kinetics of ChlZ /ChlZ formation at 80K is largely unchanged by the depletion of RC

+ carotenoids. However, the rates of formation of ChlZ at 210 K were appreciably reduced in Car-bleached PS II. Hence it was concluded that while there exist electron transfer

+ pathways from ChlZ to P680 that do not involve CarD2 at lower temperatures, the contribution of CarD2 becomes much more significant at higher (physiological) temperatures.

1.4 Implications of photosynthesis in biofuel production

Given the growing concerns about the world‘s energy demand and supply, global warming and environmental impacts of fossil fuel combustion, combined with the limited reserves of petroleum, it is necessary to develop alternative energy sources that are renewable, sustainable and carbon neutral. Bio-based fuels or biofuels that are in some way derived from biomass embrace the aforementioned caveat and therefore have the potential to replace petroleum and alleviate impending stresses on economic, environmental and energy security. In addition to being renewable and unlike petroleum based fuels, biofuels are also considered carbon neutral since they utilize/capture atmospheric CO2 in their synthesis.

Oxygenic photosynthesis plays an essential role in all biofuel production. As explained previously, it drives the conversion of light energy to chemical energy and is therefore ultimately responsible for the production of feedstocks required for all biofuels synthesis, including protons and electrons for bio-hydrogen production, sugars and starch for bio-ethanol production, oils for bio-diesel production and biomass for bio-methane production (Fig. 1.18) (reviewed in Hankamer et al., 2007). Moreover, photosystem II, equipped with its own peripheral light-harvesting complex and reaction center, is at the very center of the process of oxygenic photosynthesis and uses light energy to drive the oxidation of water (to form molecular oxygen) and production of ATP and NADPH needed for downstream processes that convert CO2 to sugar and/or oils.

Fig. 1.18 Photosynthetic biomass and biofuel production by green algae (Hankamer et al.,

2007, Physiol. Plant. 131: 10 – 21).

Biofuels such as bio-ethanol and biodiesel are presently being produced from food crops such as corn and soybean. However, from the current standpoint, these sources cannot realistically satisfy even a small fraction of the existing demand for transport fuels. Furthermore, the sole use of food crops for biofuel production is also unsustainable.

Employing microalgae as a non-food biofuel feedstock on the other hand, presents itself as a realistic and viable alternative for fulfilling growing energy demands (Dismukes et al., 2008). The potential of microalgae as a biodiesel feedstock is detailed in Table 1.2.

Microalgae can have oil contents (% oil by weight in biomass; Table 1.2, column 2) comparable to oil crops such as Camelina. However, the attainable oil yield (L oil/ ha year) is much higher for microalgae as seen in column 3 of Table 1.2. This is attributable to factors such as rapid growth rates (doubling time of ~12-24 hrs) and the possibility of harvesting fuel on daily basis rather than seasonally. Additionally, the use of microalgae as biofuel feedstock provides an advantageous scenario where 100% of the biomass may be harvested for oil. The combined effects of these benefits of algae over oil crops, result in the reduction of the land area required for cultivation and a consequent increase in biodiesel productivity (kg biodiesel/ ha year) (Table 1.2, columns 4 and 5).

Despite a host of advantages in using algae for biofuel production, the cost associated with growing and harvesting algae is still too high to make it economically feasible and competitive with petroleum. However, targeted improvements such as the use of genetically modified strains to overcome metabolic bottlenecks or sophisticated biofuel harvesting technologies would have a dramatic impact on production costs and could aid in making microalgal biofuels a pragmatic alternative to petroleum based fuels.

Moreover, increasing our understanding of the primary processes of photosynthesis in natural photosynthetic systems can abet our efforts in designing more efficient biofuel production systems.

Table 1.2 Comparison of microalgae with other biodiesel feedstocks (adapted from Mata et al., 2010, Renewable and Sustainable Energy Reviews 14: 217 – 232).

1.5 This work

Given the importance of photosynthesis and photosystem II in the production of biofuels, this thesis represents a detailed study of the Photosystem II complex including factors that govern the light-harvesting capacity of peripheral PS II antennae and the balance between forward and back electron transfer in the PS II reaction center, which are important in determining the overall efficiency of photosynthesis and biofuel/biomass production. Chapter 2 outlines genetic strategies to modulate the size of the peripheral antennae associated with PS II and their impact on photosynthetic productivity. Chapters

3 and 4 describe the effects of site-specific mutations of the PS II RC polypeptides (D1 and D2) that impact the redox properties of the primary donor (ChlD1) and terminal acceptor (QB) of PS II. The results and analysis correlate the changes in the donor and acceptor side protein environments with the efficiencies of charge separation and back electron transfer (charge recombination) in PS II and their ultimate impacts on photosynthetic productivity.

For the reasons detailed in section 1.4, the use of microalgae as a biofuel feedstock is becoming increasingly attractive. Hence the studies characterized in this thesis employ the use of the unicellular green alga Chlamydomonas reinhardtii, as a model system to study energy and electron transfer processes in PS II and their impacts on photosynthetic efficiency and overall culture productivity. Chlamydomonas reinhardtii provides a particularly good system for studying photosynthesis as Chlamydomons cells can utilize heterotrophic sources of carbon such as acetate, to grow (Harris, 1989). Hence mutants of photosynthesis can be rescued and readily grown for subsequent analyses. The nuclear

83 and chloroplast genome sequences of Chlamydomonas are also published which aids in reverse genetics studies using this alga (Merchant et al., 2007). Additionally, protocols for the efficient transformation of the nuclear (Kindle, 1990) and chloroplast (Roffey and

Sayre 1990; Roffey et al. 1991) genomes are available, making it possible to study the effects of targeted mutations of the nucleus and chloroplast. Lastly, like higher plants,

Chlamydomonas reinhardtii contains a chloroplast and hence provides a good model system for studying photosynthesis in eukaryotes.

Chapter 2

Effects of Modulating PS II Peripheral Antenna Size on Photosynthetic Productivity

and Growth of C. reinhardtii

2.1 Introduction

Algae absorb sunlight via their light harvesting or antenna complexes (LHC), which transfer excitation energy to the reaction center complexes of photosystems II (PS-II) and

I (PS-I) that drive linear electron transfer and oxygenic photosynthesis. In plants and green algae, the light harvesting antenna for PS I (termed LHCI) and PS II (termed

LHCII) bind light harvesting pigments including Chl a and Chl b and carotenoids. In nature, algal cells may acclimate to altered light environments to optimize energy capture and conversion efficiency. Cells acclimated to low light typically possess larger light harvesting antenna than those acclimated to high light intensities so as to maximize light capture at limiting light conditions. However a negative consequence of having very efficient light harvesting complexes is that photosynthetic electron transfer in nearly all photosynthetic eukaryotes reaches light saturation at only 25% of full sunlight intensity

(2000 µmol photons m-2 s-1) (Fig 2.1) (Polle et al., 2002). At high photon flux densities, the rate of photon absorption far exceeds the rate at which photosynthesis can

85 convert the excited states into charge transfer processes. Over-excitation of the light harvesting antenna under high light increases the potential for long-lived excited states and photo-oxidative damage in photosynthetic organisms due to the generation and accumulation of Chl triplets and reactive oxygen species (Krieger-Liszkay et al., 2008;

Vass and Cser, 2009). Hence algae as well as plants have both short and long term responses that protect the photosynthetic apparatus from the harmful effects of excess light (reviewed in Niyogi, 2009). Short term responses include the thermal dissipation of excess absorbed photons (qE) and state transitions (qT) both of which are components of non-photochemical quenching of chlorophyll fluorescence (NPQ). The qE (energy- dependent quenching) processes involve the de-excitation of Chl singlet excited states formed in the PS II antenna upon light absorption to minimize the formation of Chl triplets and reactive oxygen species in the photosynthetic apparatus (Müller et al., 2001).

Processes associated with qT are involved in regulating the relative excitation of PS II and PS I and help regulate the balance between linear and cyclic electron flow during photosynthesis (Wollman, 2001; Eberhard et al., 2008). Longer term responses occur over hours and days after high light exposure and include transcription and translation level changes in LHC mRNAs and high light induced mRNAs, D1 turnover and PS II repair, and increases in the xanthophyll cycle pool (Niyogi, 2009). Hence, under high light intensities, up to 80% of absorbed photons can be dissipated as heat or fluorescence due to the activation of the short term photoprotective responses (NPQ) causing large decreases in light utilization and photosynthetic productivities (Polle et al., 2002).

Fig. 2.1 Rates of photosynthesis and light absorption as a function of light intensity. This figure was adapted from Polle et al., 2002, Int. J. Hydrogen Energy 27: 1257 – 1264.

Chlamydomonas cells acclimated to high light intensities have ~50% lower cellular

Chl contents and show only slight (if any) increases in Chl a/b ratio (Neale and Melis,

1986). Most of this decrease in Chl content is associated with a reduction in PS I and

LHCI content. Additionally, increased photosynthetic electron transfer capacity is observed in high light acclimated Chlamydomonas cells on a per Chl basis (Neale and

Melis, 1986). It is not clear, however, if the same increase is observed on a per cell basis.

This is an important distinction since photosynthetic capacities calculated on a per Chl basis have an inherent bias when trying to assess changes in photosynthetic productivity.

At the molecular level, adjustment of PS II antenna size that occurs in Chlamydomonas cells acclimated to high light involves changes in nuclear LHCBM gene expression that is regulated both transcriptionally and post transcriptionally (Teramoto et al., 2002;

Durnford et al., 2003; Mussgnug et al., 2005). However the size of the PS II antenna complex is reduced by less than ~25% during the HL acclimation of Chlamydomonas

(Neale and Melis, 1986). The PS II light harvesting complex includes the proximal antenna Chl a binding proteins (CP43 and CP47) associated with the PS II reaction center; and the peripheral (distal) antenna Chl a, Chl b, and carotenoid binding proteins

(Table 2.1; Fig 2.2). The peripheral antenna complex of PS II (LHCII) further comprises of the major (outer) more abundant trimeric antenna that is encoded for by nine genes

(LHCBM1-LHCBM9) and the minor (inner) antenna proteins (CP26 and CP29) encoded for by LHCB4 and LHCB5 respectively (Fig. 2.2) (Minagawa and Takahashi, 2004; Elrad and Grossman, 2004). LHCII proteins account for binding up to 50% of the total Chl in plant and algal thylakoid membrane.

Bound Pigments Protein Chl a Chl b Chl a/b Car

Based on Guskov CP43 13 3 et al., 2009 Proximal Antenna CP47 15 5

Major Peripheral LHC-II 8 6 1.33 4 Summarized from Antenna monomer Barros and Kühlbrandt, 2009 Minor Peripheral CP29 6 2 3 ~3 Antenna CP26 6 3 2 ~3

Table 2.1 Pigment composition of the PS II proximal and peripheral antenna proteins.

Summarized based on Guskov et al., 2009 and Barros and Kühlbrandt, 2009.

Fig. 2.2 Subunit structure of PS II–LHCII supercomplex (Minagawa and Takahashi,

2004, Photosynth. Res 82: 241 – 263).

The lack of Chl b in Chlamydomonas has been shown to affect the assembly of the major peripheral light harvesting complex associated with PS II reducing the functional size of the PS II antenna from 320 Chl a and b molecules to about 95 Chl a molecules

(Polle et al., 2000). The PS I antenna, LHCI (290 Chl), and the minor LHCII complex can still assemble presumably by replacing Chl a for Chl b. Chl b is synthesized from Chl a by the action of the enzyme chlorophyllide a oxygenase (CAO) (von Wettstein et al.,

1995) and insertional mutants of Chlamydomonas that lack a functional CAO gene, lack

Chl b (Tanaka et al., 1998). Conversely, the overexpression of the CAO gene leads to the enhancement of Chl b biosynthesis in Arabidopsis and consequently to an enlargement of the PS II-associated peripheral antenna (Tanaka et al., 2001). Significantly, Chl b-less mutants (cbs-3) of Chlamydomonas have substantially elevated light-saturated photosynthetic oxygen evolution rates (up to 2.5 fold when expressed on a per Chl basis) compared to the wild-type and do not light saturate at full sunlight intensities (Polle et al.,

2000). In contrast, wild-type Chlamydomonas light saturates photosynthesis at 25% of full sunlight intensity. Moreover, studies where the size of the LHCII has been preferentially attenuated have shown that reducing PS II antenna size (and not PS I) results in higher rates of oxygen evolution at high light intensities than wild-type cells

(Polle et al., 2001, Polle et al., 2002).

We hypothesized that Chlamydomonas cells with intermediate levels of LHCII content (intermediate between wild-type and the Chl b less mutant) would maximize both light absorption and conversion at low and high light intensities. In limiting light conditions when photosynthesis is not saturated, algal cells with intermediate levels of

LHCII should prove to be more productive than small PS II antenna mutants as they have a larger PS II light absorption cross-section. On the other hand, at high light intensities, algae with intermediate levels of LHCII should be more productive than the wild-type as they have a smaller PS II light absorption cross-section allowing for more efficient light utilization presumably due to lowered energy dissipation by NPQ but are still capable of carrying out state transitions, or energy redistribution between photosystems, unlike the

Chl b less mutant. Smaller PS II antennae would also reduce mutual shading between cells and lead to better light penetration into deeper parts of the culture (Fig 2.3).

Fig. 2.3 Scenario for light absorption and utilization by algae with large and small PS II antennae at high light intensity.

Here we use an RNAi approach (Schroda, 2006) to down-regulate the expression of the CAO gene and show that the size of the PS II peripheral antenna complex assembled is directly dependent on the amount of Chl b present. Recently, RNAi technology was used to simultaneously decrease the size of LHCI and LHCII by targeting a highly conserved region of the LHC gene super family corresponding to helix 3 (Mussgnug et al., 2007). In this study, however, we pursue a ―one-gene‖ strategy to generate C. reinhardtii transformants with altered PS II peripheral antenna sizes (intermediate between wild-type and the Chl b-less mutant) by reducing the amount of Chl b within the cell. In addition, we describe a strategy for generating transgenic algae that are capable of modulating their PS II peripheral antenna size as a function of light intensity. Wild-type algae have pre-existing mechanisms to modulate the expression and size of their PS II light-harvesting antenna at the transcriptional and posttranscriptional level under varying light levels (Durnford et al., 2003). However, the range of PS II antenna adjustment in

Chlamydomonas is limiting. We take advantage of a recently described light regulated and redox-sensitive, trans-acting factor (NAB1) that binds to LHCII mRNAs, negatively regulating their translation leading to a reduction of LHCII content under high light growth conditions (Mussgnug et al., 2005, Wobbe et al., 2009). This nucleic acid binding protein 1 (NAB1) binds to a cold-shock domain consensus sequence (CSDCS) motif found in several LHCII mRNAs, sequestering them into translationally silent messenger ribonucleoprotein complexes. We have utilized the CSDCS element of the LHCMB6 mRNA to control the expression of the CAO gene in a light regulated manner. At high light intensity the NAB1 protein binds to its respective mRNA binding site on the

94 engineered CAO transcript, repressing its translation and the synthesis of Chl b, resulting in a reduced PS II peripheral antenna size. Conversely, under lower intensities translational repression by NAB1 is reduced allowing for increased levels of CAO translation and Chl b synthesis leading to the assembly of wild-type levels of the peripheral PS II antenna and in increased light capture at lower light intensities. Analogs of NAB1 have been identified in closely related algal species and hence it may be possible to extend this light-dependent post-transcriptional regulatory mechanism to Chl b synthesis in other algal species (Wobbe et al., 2009).

2.2 Materials and Methods

2.2.1 DNA constructs

The plasmid for inducing RNAi-mediated silencing of the CAO gene in

Chlamydomonas reinhardtii strain CC-424 (arg2 cw15 sr-u-2-60 mt-, Chlamydomonas

Genetic Center) was constructed using the genomic-sense/cDNA-antisense strategy of

Fuhrmann et al., 2001. A 1032 bp fragment spanning the first two exons and introns was amplified from Chlamydomonas strain CC-424 (considered wild-type for light harvesting) genomic DNA using forward and reverse primers CAOEx12GS_F and

CAOEx12GS_R (Table 2.1) respectively and cloned into the NdeI and BamHI sites of the PSL18 vector (Depege et al., 2003; a gift from Dr. Patrice Hamel, The Ohio State

University). Genomic DNA isolation from CC-424 was carried out using the xantine buffer protocol (Tillett and Neilan, 2000; http://www.chlamy.org/methods/dna.html). The corresponding cDNA region spanning exons 1 and 2 (604 bp) of CAO was then obtained

95 using RNA extracted from the CC-424 strain and cDNA synthesis (Promega). RNA extraction was performed using the manufacturer‘s protocol for trizol (Invitrogen) extraction. The 604 bp cDNA fragment was amplified using the forward and reverse primers CAOEx12CAS_F and CAOEx12CAS_R (Table 2.2), respectively and cloned in an antisense direction into the PSL18 vector using the EcoRI and BglII sites to generate the CAO-RNAi vector (Fig. 2.4). The resulting plasmid was sequenced using primers

PSL18-F-seq and PSL18-R-seq shown in Table 2.2. The PsaD promoter and terminator cassette of the PSL18 vector was used to drive RNAi. The pSL18 vector (backbone) also contains the paromomycin resistance gene driven by the Hsp70/RbcS2 fusion promoter

(Sizova et al., 2001), placed in tandem with the PsaD promoter and terminator cassette

(Depege et al., 2003). Hence transformants generated using the PSL18 vector can be selected based on resistance to paromomycin.

For the construction of the NAB1 regulated CAO, the CAO gene was amplified with the N1BSCAO-F forward and CAO-Rev reverse primers using genomic DNA isolated from Chlamydomonas strain CC-2137 (Chlamydomonas Genetic Center) as template

(Table 2.2). The 13-bp NAB1 binding site (N1BS) in the forward primer is underlined and in red. Genomic DNA was extracted from Chlamydomonas using the xantine buffer protocol as mentioned above. The NdeI and XbaI restriction sites were used in cloning of the amplified gene into the nuclear gene expression vector PSL18, to generate the PSL18-

N1BS-CAO vector (Fig. 2.3). To generate control plasmids in which the CAO gene was not preceded by the NAB1 binding site (PSL18-CAO) or had an altered NAB1 binding site (PSL18-altN1BS-CAO), the CAO-F or altN1BSCAO-F forward primers were used

96 respectively, in combination with the same reverse primer as above (Table 2.2). All the resulting plasmids PSL18-CAO, PSL18-N1BS-CAO and PSL18-altN1BS-CAO were sequenced using the PSL18-psaD-F and CAO-seq primers (Table 2.2).

Primer Sequence (5'3')

CAOEx12GS_F GCTTTCGTCATATGCTTCCTGCGTCGCTTC

CAOEx12GS_R CTCTGGATCCGTCTGTGTAAATGTGATGAAGC

CAOEx12CAS_F GACGAATTCGTCAGATGCTTCCTGCGTCG

CAOEx12CAS_R CTCTAGATCTGTCGCCTCCGCCTTCAGCTC

PSL18-F-seq CAGTCCTGTAGCTTCATACAAACATACGCACC

PSL18-R-seq GATCCTCCTGTGGCTAATTGACC

N1BSCAO-F ATCTTCATATGGGCCAGACCCCCGCAGGGCTTCCTGCGTCGCTTCAACGCAAGG

CAO-Rev TAGAATCTAGACTAGTTGTCCATGTCATCCTCGTCCACCGAG

CAO-F ATCTTCATATGCTTCCTGCGTCGCTTCAAC

altN1BSCAO-F ATCTTCATATGGGGCAAACACCGGCGGGCCTTCCTGCGTCGCTTCAACGCAAGG

PSL18-psaD-F GTTAGGTGTTGCGCTCTTGAC

CAO-Seq GGCGAGTGAGCATATTCGTCC

Table 2.2 List of primers used for cloning of the CAO-RNAi and N1BS-CAO gene constructs. The restriction sites used in cloning are in bold.

Fig. 2.4 Gene constructs used for the modulation of Chl b synthesis in Chlamydomonas.

2.2.2 Generation and screening of the CAO-RNAi transformants

For the generation of the CAO-RNAi lines (CR), the cell wall-less CC-424

Chlamydomonas strain was transformed with the CAO-RNAi plasmid by glass bead- mediated nuclear transformation (Kindle, 1990). Briefly, the CC-424 (arg2 cw15 sr-u-2-

60 mt-) cells were grown in 100 mL of Tris-Acetate-Phosphate (TAP) media (Harris,

1989) containing 100 µg/mL L-arginine (Sigma) and harvested after 4-5 days of growth

(mid-log phase) by low speed centrifugation. The cells were resuspended in 900 µL of

TAP plus 40 mM sucrose and divided equally into 3 tubes containing 300 mg 500 micron acid washed sterile glass beads. After the addition of 100 µL of 20% PEG and 1 µg of

ScaI linearized CAO-RNAi plasmid, each tube was vortexed at maximum speed for 25 sec. The cells were then resuspended in 3 mL TAP plus arginine media and grown for 24 h on a lighted incubator-shaker. Following this, the cells were spread on to TAP agar plates containing 100 µg/mL L-arginine and 50 µg/mL paromomycin for selection of the transformants. After two weeks colonies began to appear and were transfered to fresh media containing the selection antibiotics. Transformants were further screened by pigment extraction and spectrophotometric analysis of Chl a/b ratios which were expected to increase as a consequence of CAO gene silencing. For this, cells were grown in culture tubes containing 3 ml of High Salt (HS) (Harris, 1989) + Arginine (100 μg/ml) for 5-6 days under continuous illumination of ~50 μmol light m-2 s-1. The relative amounts of Chl a and b and the total Chl content was determined from 1 ml of culture

(OD750nm = 1) using the method of Arnon (1949). In addition, real-time RT-PCR analysis of selected CR transformants was done to confirm the knockdown of the CAO gene.

100

RNA from the CR transformants (CR-15, 28, 56, 68, 118 and 133) and CC-424 wild-type was extracted using Trizol according to manufacturer‘s instructions (TRI Reagent®,

Ambion, Catalog # AM9738). After treatment with DNase (Promega, Catalog # M610A),

RNA was precipitated using 3M Na Acetate and 95% ethanol. The concentration and quality of RNA was assessed using a NanoDrop spectrophotometer (Thermo Fisher

Scientific Inc.) and only samples that had 260/280 and 260/230 ratios of above 1.8 were used for further analysis. DNase-treated RNA samples (1-2 μg) were reverse transcribed with an anchored oligo (dT) primer and 200 units superscript II reverse transcriptase

(Invitrogen, Carlsbad, CA, USA) in a final reaction volume of 20 μl according to the manufacturer‘s instructions. To check whether reverse transcription (RT) and cDNA synthesis was successful, PCR was performed with the CQCBLP-F and RTCBLP-R primers (Table 2.3) using 10 μl of the RT reaction as template. Real-time quantitative

RT-PCR was carried out using an ABI – Step one plus machine using SYBR Green PCR

Master Mix Reagent Kit (Quanta Biosciences). The Chlamydomonas CBLP gene was used as reference gene/internal control and was amplified in parallel with the target gene allowing gene expression normalization and providing quantification (Zhao et al, 2008).

All reactions were carried out with 10 ng RNA in a final reaction volume of 20 µl. The primers used for the amplification of the CBLP and CAO genes are shown in Table 2.3.

The annealing temperature was set at 61 C. Each sample was set up in quadruplicates to ensure the reproducibility of the results. The quantification of the relative transcript levels was performed using the comparative CT (threshold cycle) method (Livak et al., 2001).

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Primer Sequence (5'3') CQ_CBLP_F GTGCAGGACGTGGTCATCTC

RT_CBLP_F GCAAGTACACCATTGGCGAGC

RT_CBLP_R CCTTTGCACAGCGCACAC

RT_CAO_F GACTTCCTGCCCTGGATGC

RT_CAO_R GGGTTGGACCAGTTGCTGC

Table 2.3 Primers used for real-time PCR analysis of the CAO-RNAi (CR) transformants and CC-424 (WT).

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2.2.3 Generation of NAB1 regulated CAO transformants

The PSL18-CAO, PSL18-N1BS-CAO and PSL18-altN1BS-CAO plasmids were used to generate the complemented wild-type strains (CAO), N1BS-CAO and altN1BS-CAO transgenic strains. In separate transformation events the plasmids were introduced into the CAO deletion strain cbs-3 (a gift of Dr. Ayumi Tanaka, Kyoto University) by particle gun bombardment. Briefly, the cbs-3 culture was grown in TAP media for ~5-6 d, and harvested by centrifugation while still in log phase. The concentrated cells were spotted onto the center of a TAP agar plate (~0.4-0.5 mL) and dried in a sterile-transfer hood.

Chlamydomonas nuclear transformation was carried out using a Bio-Rad PDS-1000/He system where DNA-coated gold particles were accelerated into cells of the host strain cbs-3 using pressurized helium. The gold particles (1.2 µM diameter, InBio Gold) were sterilized with ethanol and resuspended in water prior to binding of the DNA. A 50 µL aliquot of 60 mg/mL sterile gold particles was mixed with 5 μL of plasmid DNA (1 mg/mL), 50 μL of 2.5 M CaCl2, and 20 μL 0.1 M spermidine and incubated at room temperature for 20-30 min. The mixture was then centrifuged and precipitated 70% ethanol before a final resuspension in 45 µL of 95% ethanol. 15 µL of the gold coated

DNA mixture was pipetted on to a sterile macrocarrier (InBio Gold) and dried. The macrocarrier holder was installed into the chamber of the biolistics system about ~15 cm above the TAP agar plate containing the cells to be transformed and the chamber evacuated under 25 inches of Hg. The particles were then propelled into the target cells using 110 psi of helium pressure. The bombarded cells were then recovered overnight in

3 mL of TAP medium and spread onto 4-5 TAP plates containing 50 μg/mL ampicillin

103 and 25 μg/mL paromomycin. The plates were incubated at 21°C under dim light for about two weeks until small green, paromomycin resistant colonies began to appear. The colonies were transferred to fresh TAP plates containing 50 µg/mL paromomycin to minimize spontaneous transformants or escapes. Following one week of growth on TAP plates containing the selection antibiotic, DNA was extracted from the transgenics via the

Chelex-100 extraction method as described in Cao et al., 2009. Briefly, a small loop of cells was boiled in 50 µL of a 5% (w/v) solution of Chelex-100 resin (Bio-rad) for 10 min. The mixture was vortexed and spun down for 2 min to pellet the cell debris and the supernatant containing the transgenic DNA was used as the template for PCR and subsequent sequence confirmation with the pSL18-psaD-F forward and CAO-Seq reverse primers shown in Table 2.2. In addition to the complemented wild-type strains, the CC-

2137 strain (Chlamydomonas Genetics Center) served as an additional control.

2.2.4 Chl fluorescence induction measurements

Cell suspensions of the wild-type and transgenic Chlamydomonas were adjusted to yield a low Chl concentration of ~2.5 µg Chl/mL in a standard cuvette, and placed in a kinetic fluorometer (FL-3500, Photon Systems Instruments). The built-in flash Chl fluorescence induction wizard was used to run the experiment with the duration of the actinic flash set to 100 µs and the induction of Chl fluorescence during the flash was measured. The values of Chl fluorescence measured at each time point were normalized to the maximum achieved during the flash for a given sample, in order to compare the

104 percentage of closed reaction centers across samples at any given time point (% saturation).

2.2.5 Non-denaturing polyacrylamide gel electrophoresis

The CC-424 (WT), CR transformants 118 and 133, and the cbs-3 strain were grown in 100-200 mL of liquid High Salt (HS) media supplemented with 100-200 µg/mL L- arginine under low light intensities (50 µmol light m-2 s-1) with continuous shaking at 225 rpm for 6 days. Cells were harvested by centrifugation at 3,000 x g for 5 min at 4 C. The cell pellet was resuspended in buffer A (0.3M Sucrose, 25 mM HEPES, pH 7.5, 1mM

MgCl2) plus 20 µL/mL of protease inhibitor cocktail (Roche), to yield a final Chl concentration of 1 mg/mL. Cells were then broken by sonication (Biologics, Inc, Model

300 V/T Ultrasonic Homogenizer) two times for 10s each time (pulse mode, 50% duty cycle, output power 5) on ice. The unbroken cells were pelleted by centrifugation at

3,000 x g for 2 min at 4 C. The supernatant was centrifuged at 12,000 x g for 20 min and the resulting pellet was washed with buffer A. The sample was subjected to a second centrifugation step at 11,000 x g to collect thylakoids. Samples for electrophoresis were prepared by solubilization of thylakoid membranes isolated from the WT, CR-118, 133 and cbs-3 cells, with LiDodSO4 as described in Delepelaire and Chua, 1979. Briefly, the

15 µg Chl equivalent of thylakoids was solubilized in a buffer containing 50 mM

Na2CO3, 50 mM dithiothreitol, 12% sucrose and 2% lithium dodecyl sulfate to yield a final Chl concentration of 1 mg/mL and a Chl/LiDodSO4 (wt/wt) ratio of 1:20. The sample was gently shaken for 60s. Equal amounts of the sample buffer (62.5 mM Tris-

105

HCl, pH 6.8 and 25% glycerol) were added to the solubilized thylakoids before loading.

The samples were then loaded on to a Ready Tris-HCL Gel (Bio-rad 161-1225) and

LiDodSO4 and EDTA were added to the upper reservoir buffer (25 mM Tris, 192 mM glycine) to a final concentration of 0.1% and 1 mM respectively. Electrophoresis was performed at 4 ºC in the dark for 2-2.5 h at 12 mA constant current.

2.2.6 Photoautotrophic growth of CC-424, CR-118, 133 and cbs-3

The ability of the CC-424, CR and cbs-3 Chlamydomonas strains to grow photoautrophically in liquid High Salt (supplemented with 100 µg/mL L-Arginine) was measured in a time dependent manner at either low light (LL, 50 µmol light m-2 s-1) or high light (HL, 500 µmol light m-2 s-1) conditions with constant shaking at 225 rpm. The optical density of the cultures was monitored on a daily basis at 750 nm using a Cary 300

Bio UV-Vis spectrophotometer.

2.2.7 Light saturation curves of CC-424, CR-118, 133 and cbs-3

The oxygen evolving activity of log-phase cultures (0.4-0.7 OD750 nm) was assayed using a Clark-type oxygen electrode (Hansatech Instruments) (Roffey et al., 1994). Cells were resuspended in 20 mM HEPES buffer (pH 7.4) and the rate of oxygen evolution was measured as a function of increasing light intensity (650 nm red light). The photon flux density table was set to maintain the following light intensities for 1.5 minutes each: 50,

150, 300, 450, 600, 750 and 800 μE m-2 s-1 of red light, in order to obtain the light saturation curve of photosynthesis. The same experiment was repeated in the presence of

106

10 mM NaHCO3. Light saturation curves were normalized on the basis of Chl as well as cell density in order to remove any bias caused due to differences in pigment content. The

CC-424 (WT), CR-118 and CR-133 cells were grown at a light intensity of ~50 µmol light m-2 s-1. The cbs-3 strain was grown at 500 µmol light m-2 s-1, since these cells grew better at higher light intensities.

2.2.8 Pigment determination by HPLC

Chlamydomonas cultures were grown in low and high light conditions as indicated for 6 days. Aliquots of 10 ml of culture were centrifuged and the photosynthetic pigments extracted with 100% acetone in the dark for 20 min. After incubation, samples were centrifuged to pellet the cell debris and the supernatant was transferred to a glass tube and dried under vacuum. The samples were then resuspended in 750 µl of acetonitrile: water: triethylamine (900:99:1, v/v/v) for HPLC analysis. Pigment separation and chromatographic analysis was performed on a Beckman HPLC equipped with UV-vis detector, using a C18 reverse phase column at a flow rate of 1.5 mL/min. Mobile phases were (A) acetonitrile/H2O/triethylamine (900:99:1, v/v/v) and (B) ethyl acetate and the applied gradient program was 100% solvent A and 0% solvent B. The pigments were detected at 445 nm and phytoene at 282 nm.

107

2.3 Results

2.3.1 RNAi-mediated silencing of the CAO gene leads to transgenic algae with

altered Chl a/b ratios

As previously mentioned, the enzyme chlorophyllide a oxygenase or CAO is responsible for the synthesis of Chl b via the oxidation of the methyl group on ring II of

Chl a. A lack of Chl b, an abundantly found light harvesting pigment, specifically affects the assembly of the peripheral antenna complex (LHCII) associated with PS II in green algae. We used RNAi-mediated silencing to repress CAO expression and reduce cellular

Chl b levels. Here, a genomic-sense/cDNA-antisense construct spanning the first two exons and intron regions of the CAO gene was used to generate the CAO-RNAi plasmid

(Fig. 2.4). Transgenics were first identified on the basis of antibiotic resistance and colony PCR verification. Pigment analysis led to the identification of eight independent transformants (CR) that displayed a range of Chl a/b ratios between 3.2-4.9 that were 1.5-

2.2 fold higher than in the wild type (Chl a/b ratio = 2.2) with little or no change in the overall Chl content, indicating that RNAi-mediated silencing of the CAO gene leads to lower cellular Chl b levels (Table 2.4).

108

Fold Increase in Chl Total Chl per Strain Chl a/b ratio a/b ratio compared Cell (µg/mL) with WT

CC-424 (WT ) CC-424 11.3 ± 1.12 2.2 ± 0.07 1

CAO-RNAi CR-15 12.6 ± 1.76 3.2 ± 0.08 1.5

(CR transformants) CR-28 10.9 ± 1.12 3.4 ± 0.11 1.5

CR-46 11 ± 0.93 4.8 ± 0.08 2.2

CR-56 11.9 ± 0.38 3.6 ± 0.16 1.6

CR-68 11.1 ± 0.16 4.3 ± 0.33 2

CR-118 11.3 ± 0.82 3.9 ± 0.18 1.8

CR-125 10.5 ± 0.85 4.9 ± 0.20 2.2

CR-133 10.2 ± 0.51 4.9 ± 0.23 2.2

CAO-Knockout cbs-3 12.1 ± 0.98 -

(Chl-b less)

Table 2.4 Chl a/b ratios of independent CR transformants.

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2.3.2 CAO-RNAi transgenics have lower CAO mRNA levels

To determine whether the change in cellular Chl b levels and PS II antenna absorption cross section seen in the CR transformants was due to RNAi-mediated silencing of the

CAO gene, we performed real-time RT-PCR analysis using RNA extracted from the CC-

424 (WT), selected CR transformants (CR-15, 28, 56, 68, 118, and 133) and the cbs-3 mutant. The cbs-3 strain that lacks the CAO gene did not have any amplification in the real-time RT-PCR analysis and hence is not shown on the graph. On the other hand, the

CR transformants showed a 60-80% reduction in CAO mRNA levels compared to WT, confirming that the decrease in Chl b content was due to silencing of the CAO gene (Fig.

2.5).

110

Fig. 2.5 Real-time RT-PCR analysis of the CR transformants.

111

2.3.3 CAO-RNAi transgenics have truncated (intermediately-sized) PS II

antennae

To determine the effects of reduced Chl b levels on PS II antenna size in the CR transgenics, we measured the kinetics of PS II Chl fluorescence induction to its maximal level during an actinic flash (Fig. 2.6). The rate at which the Chl fluorescence rises is indicative of the rate of closure of PS II RCs and is an indirect indication of the PS II antennae size or efficiency of light harvesting (Melis, 1989; Nedbal et al., 1999). The CR transgenics had slower Chl fluorescence induction kinetics than WT and reached only 75-

85% saturation when WT reached 90% saturation. This is reflective of a smaller PS II antenna absorption cross-section in the CR mutants relative to WT. The Chl b less cbs-3 mutant did not light saturate under these measurement conditions.

112

Fig. 2.6 Chl fluorescence induction in CC-424 (WT), CR transformants and cbs-3.

113

2.3.4 Chl fluorescence rise kinetics are directly correlated with Chl b content and

size of the PS II antenna

The Chl b containing wild-type and CR strains reached their maximal Chl fluorescence values (saturation) with a single flash, in contrast to the cbs-3 strain that did not reach saturation. Arbitrarily the percentage saturation or reaction center closure was calculated for all strains at a point where wild-type algae achieved 90% saturation. When this value was plotted against the respective Chl a/b ratio, a linear inverse relationship

(with Adj. R-Square = 0.96) was observed between % saturation or RC closure and the corresponding Chl a/b ratio (Fig. 2.7). The CR transformants having higher Chl a/b ratios had slower Chl fluorescence rise kinetics presumably due to a decrease in PS II antenna optical cross-section.

To confirm a reduction in PS II antennae size in the CR transgenics, thylakoid membranes were isolated from the wild-type, two CAO-RNAi strains CR-118 and 133 and the cbs-3 mutant, and the LHC content was determined empirically using non- denaturing PAGE and densitometry (Delepelaire and Chua, 1979). We observed a ~20-

30% reduction in the intensity of the CPII band (visualized by its Chl content) corresponding to the peripheral light-harvesting antenna associated with PS II, in the CR-

118 and CR-133 transgenics indicating a decrease in LHCII content induced by CAO gene silencing (Martinson and Plumley, 1997) (Fig. 2.8). This band was absent in the cbs-3 mutant in which the assembly of the PS II peripheral antenna is affected (Fig. 2C).

The intensity of the CPI* band corresponding to the oligomeric forms of PS I was not affected in the mutants and was used as a loading control.

114

Fig. 2.7 Relationship between Chl a/b ratio and % saturation (reaction center closure).

115

Fig. 2.8 Visualization of the LHCII complex in CC-424 (WT), CR-118, 133 and cbs-3 via non-denaturing PAGE.

116

2.3.5 Photosynthetic oxygen evolution of WT, CR-118, 133 and cbs-3

It has previously been demonstrated that a reduction in the peripheral antenna size of

PS II is sufficient to increase the light saturated rate (Pmax) and capacity of photosynthesis

(Polle et al., 2000). The oxygen evolving activity of wild-type, CR-118, 133 and cbs-3 cells was measured at different intensities to yield a light-saturation curve in the presence or absence of saturating levels of CO2 added in the form of NaHCO3 (10 mM) (Fig. 2.9 and 2.10). Typically, the rate of oxygen evolution increases until the saturation irradiance is reached after which no further increase is observed. While the photon-use efficiency of photosynthesis (initial slope of the curve at low light intensity) was not impacted in the

CR transgenics, the decrease in Chl b levels (increase in Chl a/b ratios) and PS II antenna size, led to increases in the value of Pmax up to ~2 fold when compared to wild-type on a per Chl basis (Fig. 2.9). Earlier studies have demonstrated up to a 2.5-fold increase (on a per Chl basis) of Pmax in the Chl b-less cbs-3 mutant in which the PS II peripheral antenna is absent (Polle et al., 2001). Since reductions in Chl b or total Chl could give a higher photosynthetic rate measured on a Chl basis it was important to determine if photosynthesis rates also increased on a per cell basis. When compared on a per cell basis the CR transformants had up to a ~1.6 fold increase in oxygen evolution compared to WT

(Fig. 2.10). The addition of 10 mM NaHCO3 to the oxygen evolution assay showed an increase in the photosynthetic rate for all the strains. However, there was little increase in photosynthetic rate for the cbs-3 mutant in the presence of added bicarbonate possibly reflecting a reduced capacity to actively import bicarbonate. It was hypothesized that the

117 diminished ability of the cbs-3 strain to undergo state transitions had a negative impact on intracellular levels of ATP (Bulté et al., 1990; Finazzi et al., 1999; Wollman, 2001).

118

Fig. 2.9 Oxygen evolution rates of CC-424 (WT), CR-118, 133 and cbs-3 as a function of light intensity and normalized based on Chl.

119

Fig. 2.10 Oxygen evolution rates of CC-424 (WT), CR-118, 133 and cbs-3 as a function of light intensity and normalized based on cell density. 120

2.3.6 Photoautotrophic growth of WT, CR-118, 133 and cbs-3 cells

To determine if the decrease in PS II antenna size had an impact on growth rate, we measured the rate of photoautotrophic algal growth (cell density) in cultures grown at two different light intensities, 50 (LL) and 500 (HL) µmol photons m-2 s-1 (Fig. 2.11). Culture productivities of the CR transgenics were similar to the CC-424 (WT) under LL conditions, whereas the growth of the cbs-3 mutant, in which Chl b is absent, was substantially reduced having less than 50% of WT growth. The complete lack of Chl b has been shown to lower the quantum yield of photosynthesis at low light intensities

(Polle et al., 2002). Thus it is not unexpected that the cbs-3 strain grows less well (40% of

WT growth) than wild type at LL. At the HL irradiance tested here the CR strains were most productive and had a ~15-35% and 55-80% increase in growth rate compared to the

WT and cbs-3 (Chl b less) mutant respectively. Relative to growth at LL the cbs-3 strain grew at rates approaching ~75% of wild-type growth under HL.

It is interesting to note that the Chl normalized rates of O2 evolution for the Chl b less strain were higher than WT while the rates of growth were not. In green algae the assimilation of atmospheric CO2 by the Calvin cycle during oxygenic photosynthesis requires the production of ATP and NADPH in a ratio of 3:2 (reviewed in Allen, 2002).

In Chlamydomonas, the balance between linear electron flow between PS II and PS I and cyclic electron flow around PS I is in part responsible for attenuating the ATP/NADPH ratio based on cellular demands. It is possible that the ratio of linear to cyclic electron transfer in the Chl b less mutant is perturbed due to the absence of the PS II peripheral antenna. It has been shown that under physiological conditions in Chlamydomonas,

121 energetic contributions (ATP) produced as a result of state transitions as well as respiration, are required for efficient carbon assimilation and for achieving high biomass yields (Cardol et al., 2009). State transitions promote a balance between the relative excitation of PS I and PS II and almost 80% of PS II antennae are involved in this process

(Delosme et al., 1996). It is foreseeable that the lack of peripheral PS II antenna in the

Chl b less mutant could result in imbalances in linear and cyclic electron transfer and

ATP/NADPH ratios due to changes in the relative excitations of the two photosystems, in turn impacting CO2 assimilation and biomass yields. Measurements of rates of cyclic electron transfer and cellular levels of NADPH and ATP in the cbs-3 strain compared to wild-type would provide more insight into the role of PS II antenna size in determining

ATP/NADPH ratios, CO2 assimilation and biomass yields.

122

Fig. 2.11 Photoautotrophic growth of the WT, CR-118 and133 and cbs-3 cells at low and high light intensities.

123

2.3.7 PCR confirmation of the NAB1 regulated CAO transgenics

In order to achieve dynamic light-dependent modulation of Chl b synthesis and hence of PS II peripheral antenna size, we used the CSDCS element of the LHCMB6 mRNA

(NAB1 binding site or referred to here as N1BS) to control the expression of the CAO gene. At high light intensities the NAB1 protein binds to its respective mRNA binding site on the engineered CAO transcript, inhibiting its translation and the synthesis of Chl b, resulting in a reduced PS II peripheral antenna size. Under lower intensities translational repression by NAB1 decreases and wild-type levels of the peripheral PS II antenna can assemble increasing light capture at lower light intensities. A gene construct in which the

13-bp LHCBM6 mRNA CDSCS or N1BS was placed at the 5‘ end of the CAO gene, was introduced into the cbs-3 mutant by particle gun bombardment to generate the N1BS-

CAO transgenics. As a control we complemented the cbs-3 mutant with the wild-type

CAO gene lacking a 5‘ N1BS to generate the complemented wild-type. We also inserted a mutagenized NAB1 binding site (different from LHCBM6 mRNA CDSCS by 4 bp) into the 5‘ coding sequence of the CAO gene and introduced this into the Chl b-less strain, cbs-3, to generate the altN1BS-CAO transgenics. In all cases, the PsaD promoter was used drive the expression of the gene construct so as to decouple the effects of the native

CAO promoter. The PsaD promoter used in combination with its native terminator has been shown to drive constitutive and high levels of gene expression (Kumar and Sayre,

2010, manuscript in preparation). The transgenics were selected initially on the basis of antibiotic resistance and further screened by pigment extraction and quantification.

Selected transgenics having Chl a/b ratios most similar to wild-type (CC-2137) were

124 confirmed for the presence of the transgene by PCR (Fig. 2.12). The amplified region was verified via DNA sequence analysis.

125

Fig. 2.12 PCR confirmation of the transgenics containing the NAB1 regulated CAO gene constructs.

126

2.3.8 Chl a/b ratios of the N1BS-CAO transgenics

To analyze the effect of the 5‘ NAB1 binding site on the regulation of the CAO gene and Chl b synthesis during photoacclimation, the Chl a/b ratios of the individual transformants were determined by pigment extraction and HPLC analysis of cultures grown at LL (50 µmol photons m-2 s-1) or at HL (500 µmol photons m-2 s-1) for 6 days.

Each strain was inoculated into fresh HS media using a 2% (v/v) culture inoculum to avoid self-shading and nutrient limitation. Chl a/b ratios were then monitored through two sets of alternating periods of low and high irradiance as shown in Fig. 2.13. The complemented wild-type (CAO-4, 22) and CC-2137 strains showed similar trends with slight reductions (<2-6%) in Chl a/b ratios when grown under HL probably due to the effects of photoinhibition (Harper et al., 2004). The N1BSCAO-4, 7 and 77 transgenics on the other hand showed the opposite trend with up to a ~16% increase in Chl a/b ratios when grown under HL conditions. This indicated a preferential decrease in Chl b synthesis in response to high irradiances due to NAB1 regulation of CAO. Moreover, the altN1BS-CAO transgenics showed trends similar to the complemented wild-type and CC-

2137 strains where a decrease in Chl a/b ratios was observed under high light intensities presumably due to photoinhition as mentioned above. This indicated that NAB1 binding to the CAO transcript in the altN1BS-CAO transgenics was probably perturbed due to 4 bp change in the sequence of altered N1BS compared to the original NAB1 binding site.

127

Fig. 2.13 Changes in Chl a/b ratios in the complemented WT (CAO-4, 22), CC-2137

(WT), N1BSCAO and altN1BSCAO transgenics during acclimation to low and high light.

128

2.3.9 Chl fluorescence induction in the N1BS-CAO transgenics

To correlate the changes in Chl a/b ratio to possible alterations in PS II antenna size, the cells were subjected to flash Chl fluorescence induction as described before, after each light period. Percentage light saturation or reaction center closure was calculated for the transformants at a time point where the complemented wild type strain, CAO-4, achieved 90% saturation. The values obtained for each strain under low and high light were compared to yield a percentage decrease/increase in Chl fluorescence yield.

Reversible changes in Chl fluorescence induction kinetics of up to ~10% were observed after each light cycle in the N1BS-CAO transgenics as compared to less than a ~1-2% change in the CC-2137 wild-type control (Fig. 2.14).

The significant increases in Chl a/b ratios and decreases in Chl fluorescence induction kinetics in the N1BS-CAO transgenics upon being transferred from low light to high light intensities, indicated that Chl b synthesis and therefore the PS II absorption cross-section decreased under high light to a degree greater than in wild-type cells. In our experiments,

Chlamydomonas cells were light acclimated for six day periods during which the NAB1- regulated (light-dependent) changes in Chl b accumulation and PS II antennae size occurred. Hence this strategy would be applicable in maximizing photon absorption and utilization during seasonal changes in light intensity.

129

Fig. 2.14 Changes in Chl fluorescence induction in the complemented WT (CAO-4, 22),

CC-2137 (WT), N1BSCAO and altN1BSCAO transgenics during acclimation to low and high light.

130

2.4 Discussion

In this work, we demonstrate that modulating the levels of Chl b, which binds preferentially to the PS II peripheral antenna, is sufficient to affect the size of the PS II-

LHCII antenna complex but also to substantially alter biomass yields (Figs. 2.6, 2.8 and

2.11). We also confirm a linear but inverse relationship between the Chl a/b ratio and the size of the PS II peripheral antenna (Fig. 2.7). This has previously been shown to be the case in the Su/su and Su/su var. mutants of tobacco in which the yellow-green

(intermediate Chl b levels) and yellow (low Chl b) leaf sections showed increasing photosynthetic capacities with decreasing levels of Chl b (Homann and Schmid, 1967).

Here, two approaches were adopted to modify Chl b accumulation and PS II antenna size, a) the permanent truncation of the PS II antenna through RNAi mediated silencing of the

Chl b synthesis gene, CAO, and b) the dynamic and irradiance-dependent adjustment of

PS II antenna size through the post-transcriptional regulation of CAO by the light- responsive translational inhibitor protein, NAB1 (Fig. 2.4).

A number of strategies have previously been used to modulate light harvesting antenna size (Nakajima and Ueda, 1999; Polle et al., 2000; Tanaka et al., 2001; Polle et al., 2002; Polle et al., 2003; Mussgnug et al., 2007; Tetali et al., 2007; Wobbe et al.,

2009; Beckmann et al., 2009). The overwhelming consensus that emerges is that small antenna mutants have increased light utilization efficiency since they do not saturate rate- limiting, downstream electron transfer processes. However, few studies have shown how this manifests into growth under photoautotrophic conditions at low and high light intensities. As has previously been shown, organisms with smaller PS II light harvesting

131 antennae are less efficient at harvesting photons at low light intensities but do not light saturate electron transfer at high light intensities (Polle, et al., 2002; Figs. 2.9 and 2.10).

From an evolutionary perspective organisms that occupy potentially diverse light intensity regimes, like algae, are better capable of surviving under very low light conditions at great depths in the water column if they have larger antennae (Schenk et al.,

2008). The trade off is reduced efficiencies at high light intensities when electron transfer reactions are light saturated. As shown here, the CR transgenics with intermediate antennae sizes outperform wild-type cells in monocultures at light saturating intensities without an impairment in effective photosynthesis or growth rates at sub-saturating light intensities (Fig. 2.11). Although Chl b levels in the CR transgenics are lower than in WT, the total Chl content is comparable (Table 2.4) suggesting that there are higher levels of

Chl a or possibly reaction centers present. Intermediate PS II antennae in the CR transformants allow for sufficient photon capture and presumably do not disrupt the optimal ATP/NADPH ratios required for CO2 assimilation under low light conditions.

Under the high light intensities tested, the decreased PS II antennae size and increased light saturation rates of the CR transformants compared to the WT along with a possible increase in number of reaction centers leads to a faster growth rate than the WT.

Over the course of a year, light intensities and duration in the temperate regions of the world vary dramatically. It has been hypothesized that dynamic modulation of antennae size can optimize light harvesting efficiency at low and high light intensities. In separate studies, we show that cells with large antennae yield more biomass when grown in shallow ponds (8 cm) versus growth in deeper ponds (28 cm) due to self shading effects

132 of large antennae size (Stroff, Perrine, Sayre manuscript in prep). In contrast, algae with small antennae sizes produce 30% more biomass when grown in deeper ponds due to less self-shading. In ponds of constant depth it would be ideal if algal cells could self modulate their PS II light-harvesting antenna size seasonally to optimize light capture with minimal self-shading and minimal light saturation of photosynthetic electron transfer. In certain green algae, such as Dunaliella salina CAO mRNA levels change dramatically in an irradiance dependent manner decreasing up to 5-fold under high light and resulting in an increase in Chl a/b ratios from ~4 to 12 (Masuda et al., 2002; Masuda et al., 2003). In contrast, in higher plants such as A. thaliana, this light dependent regulation of CAO mRNA levels is less dynamic. A minor increase in the Chl a/b ratio occurs in Arabidopsis during high light acclimation, accompanied by a 2-fold decrease in

CAO transcript levels (Walters and Horton, 1994; Harper et al., 2004). Similarly, light adaptation responses in C. reinhardtii have revealed only slight increases in Chl a/b ratio during high light adaptation (Neale and Melis, 1986; Durnford et al., 2003). In contrast to the aforementioned studies, the photoacclimation experiments described in this study were carried out for a longer time interval (6 days) and small but consistent decreases in the Chl a/b ratios of the C. reinhardtii CC-2137 and the CAO-4, 22 complemented wild- type strains were observed under high light intensities (Fig. 2.13). A similar scenario has been observed in A. thaliana during photoinhibitory light treatment (Harper et al., 2004).

Regulation of LHCII expression in Chlamydomonas has, however, been shown to occur at the transcriptional and post-transcriptional level (Teramoto et al., 2002; Durnford et al.,

2003; Mussgnug et al., 2005). More recently, the post transcriptional regulation of LHC

133 mRNAs has been shown to involve a cytosolic translation repressor protein, NAB1, which binds to its target mRNAs via a 13 bp NAB1 binding site. High irradiances result in a reducing cytosolic environment and in the activation of NAB1 which then binds to

LHCII transcripts and repress translation decreasing Chl b synthesis and reducing PS II peripheral antenna size. This NAB1 binding activity is redox-controlled and regulated via two cysteine residues within its C-terminal RNA Recognition Motif (Wobbe et al., 2009).

The mutagenesis of the cysteines to serine residues produces a permanently active form of the protein, causing an irradiance-independent and irreversible reduction of ~10-20% in the size of the PS II antenna, accompanied by a small increase in Chl a/b ratio of ~6%

(Beckmann et al., 2009). This minor decrease in antenna size was seen as being sufficient to produce a ~30% increase in photoautotrophic growth at high light intensities.

We demonstrate here that modulation of the PS II peripheral antennae size by light- regulated control of Chl b accumulation may provide a strategy to optimize PS II antennae size in response to seasonal changes in light intensity. Using a 13 bp NAB1 binding site fused to the 5‘ end of the CAO transcript, we demonstrate that it is possible to post-transcriptionally regulate Chl b accumulation and the size of the PS II antenna complex. Our results from three independent transformants expressing the modified CAO gene show that Chl a/b ratios increase under HL acclimation indicating a reduced PS II antenna size (Fig. 2.13). Conversely, a decrease in Chl a/b ratios under conditions of low irradiances are indicative of an increase in PS II antenna size. This interpretation is supported by the observation that flash Chl fluorescence induction kinetics of low light grown cultures of the NAB1-CAO transgenics exhibited up to a ~10% increase in relative

134

Chl fluorescence yield compared to the high light grown cultures, whereas a 1-2% increase was observed in the CC-2137 and complemented wild-type strains (Fig. 2.14).

This light-dependent change in antennae size in the transgenics is substantially greater than that observed in wild-type cells. Currently, we are assessing the impact of the changes on biomass yield at various pond depths over the course of a year.

Engineering approaches have also been used to optimize light capture and energy conversion efficiency. For instance, photo-bioreactors have been developed to distribute sunlight over larger surface areas thereby reducing saturating light intensities for photosynthesis (Carlozzi, 2003). Optimizing culture mixing has also been used to reduce the likelihood of light saturation of photosynthesis (Meiser et al., 2004; Yoshimoto et al.,

2005). At the present time it is not clear if the increases in yield potentially achieved in closed photobioreactors (PBR) relative to open ponds justifies the capital expenses of

PBRs. Open culture ponds are more economically feasible in terms of construction and maintenance and the vast bulk of microalgae today are cultivated in this manner.

To provide the greatest flexibility to microalgal production systems, algal strains that self-modulate and optimize their antennae size would require minimal infrastructure investment to optimize light harvesting and energy conversion efficiency.

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Chapter 3

Site-Directed Mutagenesis of D1-T179 Modifies the Redox Properties of the Primary

Electron Donor in Photosystem II

3.1 Introduction

Photosystem II (PS II) in plants and green algae is an integral membrane pigment- protein complex that catalyzes the light-driven oxidation of water and reduction of plastoquinone. Primary charge separation takes place in the reaction center (RC) of PS II which consists of the D1, D2, cytochrome b559, and psbI proteins as well as six chlorophylls (PD1, PD2, ChlD1, ChlD2, ChlZD1, ChlZD2) and two pheophytin (PheoD1 and

PheoD2) electron transfer (ET) cofactors (Ferreira et al., 2004; Loll et al., 2005; Guskov et al., 2009). The structural organization of the core pigments in PS II RCs is similar to that in the purple bacterial reaction center (BRC). Both RC types have two parallel, C2 symmetry-related cofactor branches that span the membrane. However, only one of the two branches (active branch) participates in primary electron transfer. In the PS II RC,

* ET from the excited state of the primary electron donor (P680 ) to a Pheo molecule

(PheoD1) represents the primary photochemical event of photosynthesis and results in the

+ - formation of the primary radical pair (P680 PheoD1 ). The electron from the reduced Pheo

136 is then transferred to a bound plastoquinone molecule (QA) forming the first stable charge

+ - separated state, P680 QA . The subsequent transfer of electrons from QA to the plastoquinone molecule bound in the QB site (QB) represents the rate limiting step of ET in PS II. Unlike QA, QB accepts two electrons and undergoes protonation, after which it is released from PS II into the membrane matrix to transfer electrons to the cytochrome b6f

+ complex. The primary electron donor cation P680 is re-reduced by a redox-active tyrosine, known as TyrZ (Y161 of the D1 subunit), to generate a neutral tyrosine radical

• TyrZ . This neutral radical then acts as an oxidant for the water oxidation process that occurs at the oxygen evolving complex (OEC). The catalysis of water oxidation proceeds via the S-state cycle in which the OEC transitions through different oxidation states (S0-

S4) to produce molecular oxygen (reviewed in Ke, 2001; Kern and Renger, 2007;

Gokhale and Sayre, 2009).

There is substantial sequence as well as structural homology between the purple BRC and PS II RC (Diesenhofer and Michel, 1989; Ruffle et al., 1992). One such similarity between the two is the presence of the conserved histidine residues D1-H198 and D1-

H197 in PS II which act as axial ligands for the PD1 and PD2 Chls of the Chl special pair

(ChlSP). Despite the obvious homology between the two RCs, there are several important differences: (a) the primary donor cation generated upon excitation of PS II has a much higher oxidizing potential (~1.2 V) when compared to that of the BRC (~0.4 V) (Allen and Williams, 1998; Kato et al., 2009) (b) the exciton coupling between the PD1 and PD2

Chls of the ChlSP in PS II is far weaker than that associated with the bacteriochlorophyll special pair (BChlSP) in the BRC, most likely due to increased separation of PD1 and PD2

137 compared to their bacterial counterparts (Tetenkin et al., 1989; Braun et al., 1990;

Durrant et al., 1995; Ferreira et al., 2004) and (c) the Chl triplet formed upon charge recombination in PS II is localized on a monomeric Chl oriented 30 relative to the normal of the membrane plane while in the BRC it is localized on the BChlSP (van

Meighem et al., 1991; van Gorkom and Schelvis, 1993). Furthermore, a greater degree of asymmetry exists between the axial ligands for the RC Chls in PS II when compared to those in the BRC. More specifically, in the BRC, all the axial ligands to the RC BChls are histidine residues. On the other hand in the PS II RC, while the axial ligands of the PD1,

PD2, ChlZD1 and ChlZD2 are histidines, the active and inactive branch monomeric Chls,

ChlD1 and ChlD2, lack protein ligands. Instead, the amino acid residues D1-T179 and D2-

I178 are found to be in closest proximity to ChlD1 and ChlD2, respectively (Ferreira et al.,

2004).

In the BRC, the primary electron donor is the BChlSP, a dimer of BChl molecules

(Knapp et al., 1985, Michel et al., 1986; reviewed in Kirmaier and Holten, 1987 and

Allen and Williams, 1998). The identity of the primary electron donor in PS II, P680, however, has been the subject of considerable debate (reviewed in van Gorkom and

Schelvis, 1993; Diner, 2001; Schlodder et al., 2008a). There is an increasing body of evidence suggesting that P680 has monomeric properties including: (1) the lack of a significant red shift in the Qy absorption band as expected for an energetically coupled

Chl dimer (Tetenkin, 1989) (2) results from Stark effect and hole burning experiments that are not as expected for a Chl dimer and (3) spectroscopic evidence for the localization of a Chl triplet on a monomeric Chl in the PS II RC, probably ChlD1 (van

138

Meighem et al., 1991; van Gorkom and Schelvis, 1993; Schlodder et al., 2008a). More recently, it has been shown that the active branch monomeric Chl, ChlD1, and not the

ChlSP, has the lowest site energy and hence the excitation energy is more localized on this

RC Chl, particularly at liquid helium temperatures (Raszewski et al., 2005; Schlodder et

* * al., 2008a). This makes ChlD1 , rather than ChlSP , the plausible primary electron donor and the site of initiation of primary charge separation in PS II. At ambient temperatures, however, the excited state is delocalized over the entire multimeric set of RC chlorins that include the ChlSP, ChlD1/D2, and PheoD1/D2, but primary charge separation is still initiated from ChlD1 (Durrant et al., 1995; Dekker and van Grondelle, 2000; Barter et al.,

+ - 3 1 2003). Flash induced (P680 QA —P680QA) and ( P680- P680) absorbance difference spectra of the D1-H198Q and D1-T179H Synechocystis mutants with respect to WT, have

3 + - indicated that the RC triplet that arises from P680 PheoD1 recombination is localized on

ChlD1 whereas the oxidized primary donor cation is localized on the PD1 Chl of the PS II

ChlSP (Schlodder et al., 2008b). In addition, recent reports have found that the reduction of PheoD1 occurs prior to the oxidation of PD1/PD2 of the ChlSP, even at ambient temperatures, implying that the cation of the primary radical pair is localized at a site other than the ChlSP, most likely ChlD1 (Groot et al., 2005 and Holzwarth et al., 2006).

The efficiency and yield of primary charge separation in PS II is dependent on the ability of the RC to generate and stabilize charge separated states, which is, in part, influenced by the redox potentials and energetic gaps between donors and acceptors.

Analyses of site-directed mutagenesis of PS II residues such as D1-E130 and D1-H198 have demonstrated that the local protein environment plays an important role in

139 influencing the redox potentials of ET cofactors in PS II (Roffey et al., 1994; Giorgi et al., 1996; Dorlet et al., 2001; Rappaport et al., 2005; Cser K and Vass I., 2007). Moreover computational studies using the linearized Poisson-Boltzmann equation and the crystal structure of PS II at 3.2 Å resolution, have shown that the D1 and D2 RC proteins contribute significantly to the redox potentials of the RC Chls (PD1, PD1, ChlD1, ChlD2), accounting for the sizable increase in redox potential from ~+0.8 V for Chl a in dimethylformamide (DMF) to ~+1.1 - 1.3 V for Chl a in the PS II RC core (Ishikita et al.,

2005). In particular, the axial ligands of the ChlSP D1-H198 and D2-H197 were predicted as influencing the redox potentials of the PD1 and PD2 Chls respectively, while the D2-

R180 residue was predicted as contributing to the redox potential of ChlD2. However, no specific amino acid residue in the vicinity of ChlD1 was predicted as having an effect on the redox potential of ChlD1.

Given that ChlD1 is the most probable candidate for being the primary electron donor in PS II, we modified its local protein environment to elucidate the function of ChlD1 in charge separation as well as to study the effects of the proximal amino acid residues on its redox properties. D1-T179 is the closest residue to ChlD1 and overlies the coordinated

2+ Mg but does not serve as a direct ligand (Fig. 3.1). Axial coordination to ChlD1 is probably provided by a water molecule hydrogen bonded to D1-T179O. Despite not being involved in directly ligating ChlD1, D1-T179 is completely conserved throughout oxygenic photosynthesis. We performed site-directed mutagenesis of the psbA gene in

Chlamydomonas reinhardtii in which D1-T179 was replaced by serine, aspartate, asparagine, histidine and isoleucine. The resulting Chlamydomonas mutants were

140 analyzed using flash-induced Chl fluorescence decay kinetics and thermoluminescence

(TL). We show that mutagenesis of this residue impairs photoautotrophic growth and photosynthetic oxygen evolution. Furthermore, the substitution of D1-T179 to more polar residues such as aspartate, histidine and asparagine (Radzicka and Wolfenden, 1988) increases the free energy level of the primary radical pair and results in accelerated rates of charge recombination. The opposite effect is seen when the substituted residue is isoleucine. The replacement of D1-T179 with isoleucine and asparagine also led to an increase in basal Chl fluorescence levels suggesting that D1-T179 functions in mediating excitation energy transfer from the proximal PS II antenna to the RC complex.

Interestingly, the D1-T179I mutant showed the lowest levels of photoautotrophic growth and oxygen evolution. It is noteworthy that the inactive branch residue that corresponds to D1-T179 is also an isoleucine (D2-I178) (Ferreira et al., 2004).

Our results show that the nature of the residue in position 179 of the D1 RC- polypeptide (closest to ChlD1) is involved in mediating excitation energy transfer to the

RC and is paramount in determining the redox properties of the primary electron donor.

The latter plays a critical role in balancing forward ET with charge recombination.

141

Fig. 3.1 The protein environment of ChlD1 and the relative location of D1-T179. The figure was based on Protein Data Bank (PDB) coordinates 3BZ1 (Guskov et al., 2009) and was generated using PyMOL.

142

3.2 Materials and Methods

3.2.1 Bioinformatics analysis

Sequences for the Photosynthetic Reaction Center Apo-Protein family (pfam00124) and the PS II RC D1 protein family (CHL00003) were obtained from the Conserved

Domain Database, CDD, (http://www.ncbi.nlm.nih.gov/Structure/cdd/cdd.shtml) and a

Position-Specific Scoring Matrix (PSSM) was generated for the photosynthetic reaction center and the D1 protein families through the ―Statistics‖ tab. The PSSM of a protein family contains the log-odds score or residue frequency for finding a particular amino acid in a given position of the consensus sequence and is calculated based on a multiple alignment of all the sequences within that family of proteins. Hence it yields information about the relative conservation of individual amino acid residues within protein families and facilitates the identification of potential functionally important residues. Residue frequencies for positions 132 and 83, corresponding to the protein environment surrounding the monomeric RC Chls and Pheos respectively, of the consensus sequence of all type II RC polypeptides, were obtained. Similarly, residue frequencies for positions

179 and 130 of the consensus sequence of the D1 protein family were acquired. The residue frequencies for the two protein families were then compared to reveal critical differences in the protein environment of the Chl (BChl) monomers and Pheos (BPheos) of the two cofactor branches in anoxygenic (bacterial RCs) and oxygenic (PS II RCs) photosynthetic reaction centers.

143

3.2.2 DNA constructs

Five different point mutations were introduced at position 179 of the D1 protein in which threonine (wild-type) was replaced with isoleucine (D1-T179I), serine (D1-

T179S), aspartate (D1-T179D), histidine (D1-T179H) and asparagine (D1-T179N). The intron-less version of the Chlamydomonas reinhardtii psbA gene (encodes the D1 protein) present in the pBA155 plasmid was used as the template for site-directed mutagenesis. The map of the parent plasmid, pBA155, is described in Minagawa and

Crofts, 1994. The QuikChange site-directed mutagenesis kit from Stratagene along with the primers pairs shown in Table 3.1 was used to create the mutagenized plasmids. The plasmids were confirmed as possessing the desired mutations by PCR and sequencing.

144

Primer Sequence (5'3')

D1-T179S-F GCCTTTAGGTATCTCTGGTAGTTTCAACTTCATGATCGTATTCC

D1-T179S-R GGAATACGATCATGAAGTTGAAACTACCAGAGATACCTAAAGGC

D1-T179D-F GCCTTTAGGTATCTCTGGTGATTTCAACTTCATGATCGTATTCC

D1-T179D-R GGAATACGATCATGAAGTTGAAATCACCAGAGATACCTAAAGGC

D1-T179H-F GCCTTTAGGTATCTCTGGTCATTTCAACTTCATGATCG

D1-T179H-R CGATCATGAAGTTGAAATGACCAGAGATACCTAAAGGC

D1-T179N-F GCCTTTAGGTATCTCTGGTAATTTCAACTTCATGATCGTATTCC

D1-T179N-R GGAATACGATCATGAAGTTGAAATTACCAGAGATACCTAAAGGC

D1-T179I-F GCCTTTAGGTATCTCTGGTATTTTCAACTTCATGATCGTATTCC

D1-T179I-R CGATCATGAAGTTGAAAATACCAGAGATACCTAAAGGC

D1-5'UTR GGACGTAGGTACATAAATGTGCTAGGTAAC

D1-3'UTR CCTGCCAACTGCCTATGGTAGCTATTAAGT

Table 3.1 List of primers used for site-directed mutagenesis of D1-T179.

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3.2.3 C. reinhardtii chloroplast transformation

Plasmids containing the mutagenized psbA genes were introduced, in separate transformation events, into the psbA deletion strain CC-4147 (Chlamydomonas genetics center, Duke University) by particle gun bombardement. Briefly, the CC-4147 culture was grown in Tris-Acetate-Phosphate (TAP) media (Harris, 1989) for ~5-6 d, and harvested by centrifugation while still in log phase. The concentrated cells were spotted onto the center of a TAP agar plate (~0.4-0.5 mL) and dried in a sterile-transfer hood.

Chlamydomonas transformation was carried out using a Bio-Rad PDS-1000/He system where DNA-coated gold particles were accelerated into cells of the host strain CC-4147 using pressurized helium. The gold particles (1.2 µM diameter, InBio Gold) were sterilized with ethanol and resuspended in water prior to binding of the DNA. A 50 µL aliquot of 60 mg/mL sterile gold particles was mixed with 5 μL of plasmid DNA (1 mg/mL), 50 μL of 2.5 M CaCl2, and 20 μL 0.1 M spermidine and incubated at room temperature for 20-30 min. The mixture was then centrifuged and precipitated 70% ethanol before a final resuspension in 45 µL of 95% ethanol. 15 µL of the gold coated

TAP medium, and spread onto 4-5 TAP plates containing 50 μg/mL ampicillin and 100

μg/mL spectinomycin. The plates were incubated at 21°C under dim light until small

146 green, spectinomycin resistant colonies began to appear (usually about two weeks). The colonies were transferred to fresh TAP plates containing 100 µg/mL streptomycin

(resistance to which is also conferred by the aadA gene of the pBA155 plasmid) to minimize spontaneous transformants or escapes (Goldschmidt-Clermont, 1991).

Following one week of growth on TAP plates containing the selection antibiotic, DNA was extracted from the transgenics via the Chelex-100 extraction method as described in

Cao et al., 2009. Briefly, a small loop of cells was boiled in 50 µL of a 5% (w/v) solution of Chelex-100 resin (Bio-rad) for 10 min. The mixture was vortexed and spun down for 2 min to pellet the cell debris and the supernatant containing the transgenic DNA was used as the template for PCR and subsequent sequence confirmation with the D1-5‘ and 3‘

UTR primers shown in Table 3.1. The CC-3946 strain (Chlamydomonas Genetics Center) containing an intronless psbA gene, was used as a control in this study and is referred to as the wild-type (WT).

3.2.4 Growth of Chlamydomonas WT and D1-T179 mutants

All strains were maintained on TAP agar plates containing 50 µg/mL ampicillin to prevent bacterial contamination. In addition, plates containing the D1-T179 mutants also contained 100 µg/mL spectinomycin. For subsequent measurements, cells were grown up to mid to late log phase (4-5 days) under 25 µMol photons m-2 s-1 illumination and 225 rpm shaking in liquid TAP media in the absence of spectinomycin. The photosynthetic competence of the wild-type and D1-T179 mutants was assayed by observing their growth on photoautotrophic high salt (HS) agar media (Harris, 1989). For this, the optical

147 density of TAP-grown mid-log phase cells was measured at 750 nm using a Cary 300 Bio

UV-Vis spectrophotometer. An equal number of cells from each strain were washed and resuspended in high salt (HS) medium, diluted serially and pipetted (10 µL) on to HS agar plates and incubated at 21 C under 50 µMol photons m-2 s-1 continuous illumination. The growth of the cultures after two weeks was imaged.

3.2.5 Chl analysis

The Chl concentration of the culture was determined by vortexing the cell pellet obtained from 1 mL of culture with 80% acetone. The acetone extract was then centrifuged at 14,000 x g to pellet the cell debris. After ensuring that the pellet was white in color (indicating complete extraction), the optical absorption of supernatant was measured at 663 and 645 nm. The Chl content was calculated according to the method of

Arnon (1949).

3.2.6 Assaying oxygen evolution

The oxygen evolution activity of the wild-type and mutant cells was measured in

TAP medium under continuous illumination (~800 μMol photons m-2 s-1 of red light) in the presence of 2 mM potassium ferricyanide and 0.5 mM of 2,6-dimethyl-p- benzoquinone (DMBQ), using a Clark-type oxygen electrode from Hansatech

Instruments (Roffey et al., 1994; Rose et al., 2008). The Chl concentration of the sample was ~15 µg/mL. Final measurements were an average of six assays from at least three independent cultures for each strain.

148

3.2.7 Flash-induced Chl fluorescence measurements

- The reoxidation of QA was assayed in the WT and D1-T179 mutant cells in the presence and absence of 20 μM DCMU by monitoring Chl fluorescence decay kinetics following a single saturating flash using a pulse modulated fluorometer (Photon Systems

Instruments) in the 100 μs–100 s time range. The sample was dark adapted for 10 min prior to the flash and the Chl concentration used was 5 µg/mL. Chl fluorescence decay curves were resolved using a fitting function with three lifetime components as described in Vass et al., 1999. The fast and middle phases were simulated with exponential

- components. The slow component representative of the reoxidation of QA via charge recombination has been shown to follow hyperbolic kinetics (Bennoun, 1994). Therefore, a hyperbolic component was used to model the slow phase.

F(t) = A1exp(-t/T1) + A2exp(-t/T2) + A3/(1+t/T3) where F(t) is the raw fluorescence yield at time t, A1–A3 are the amplitudes, T1–T3 are the lifetime components.

In addition, the true values of F0 were measured prior to the actinic flash, using low intensities of measuring light to ascertain whether any increases were evident in basal levels of PS II Chl fluorescence emanating from the antenna, in response to inefficacies in excitation energy transfer to the reaction center (Xiong et al., 1997).

149

3.2.8 Thermoluminescence measurements

Thermoluminescence from whole cells was measured in the presence and absence of

20 μM DCMU using a thermoluminescence instrument manufactured by Photon Systems

Instruments. A cell suspension of 50 µg Chl per 50 µL of 20 mM HEPES at pH 7.5 and

0.5 µM Nigericin was spotted on a ½ inch disc of Whatman filter paper 1. The sample was dark-adapted for 2 min and then cooled to 0 ºC (or -10 C when measuring TL in the presence of DCMU). After 2 min, a single saturating flash was fired and TL was recorded upon heating the sample at a rate of 0.5 ºC/s.

3.3 Results

3.3.1 Protein environment of the monomeric Chls in type II RCs

The identity of the amino acid residues that serve as axial ligands to the central Mg2+ atoms of the RC Chl monomers in the BRC and PS II RC were compared using a

Position Specific Scoring Matrix (PSSM) based on the protein sequences of the

Photosynthetic Reaction Center Apo-Protein family (pfam00124) and the PS II D1 protein family (CHL00003). The residue frequencies at positions 132 and 83 of the pfam00124 consensus sequence, corresponding to positions D1-179 (Threonine) and D1-

130 (Glutamate) in Chlamydomonas reinhardtii were obtained (Table 3.2). Similarly, the frequencies of residues occurring at positions 179 and 130 of the D1 protein family consensus sequence were scored. According to the PSSMs, histidine, threonine and isoleucine are the amino acid residues most frequently found immediately adjacent to the

RC Chl monomers of type II RCs, whereas glutamate and glutamine most frequently

150 occur in the environment around the Pheo head groups (Table 3.2). In comparing the two

PSSMs for the type II RC protein family and the PS II D1 RC protein family, it becomes apparent that while the acceptor sides of the BRC and PS II RC are highly similar, the protein environment of the monomeric Chls (ChlD1 and ChlD2) on the donor side is drastically different in that histidine residues no longer serves as direct ligands to the central Mg2+ atoms. Instead, a threonine residue is in closest proximity to the Chl monomer (ChlD1) of the active branch in the PS II RC, and is found as being completely conserved, occurring in 138 (out of 139) members of the D1 protein family. Hence this residue, despite not being a direct ligand, is probably required for optimum functioning of

PS II charge separation.

151

Table 3.2 Position Specific Scoring Matrices (PSSMs) generated for the Pfam00124 and

CHL00003 protein families.

152

3.3.2 Growth characteristics of site-directed mutants of D1-T179

As previously discussed, the D1-T179 residue is the closest amino acid residue near the active branch chlorophyll monomer, in which axial coordination is probably provided by a water molecule that is hydrogen bonded to D1-T179Oγ (Ferreira et al., 2004, Loll et al., 2005). In order to gain greater insight into the structure/function role of this highly conserved residue, we carried out site-directed mutagenesis to generate the D1-T179 S,

D, H, N and I mutants. To determine whether these mutants retained their photosynthetic competencies the growth of the cells on photoautotrophic growth media was assayed.

Cells were diluted to equal optical densities and spotted on both Tris Acetate Phosphate

(TAP) (control) as well as High Salt (HS) agar plates. While all the mutants had significant growth defects, the D1-T179N and D1-T179I substitutions had the most drastic phenotypes whereas the growth of the D1-T179S mutant was the least affected

(Fig. 3.2).

153

Fig. 3.2. The growth characteristics of the WT and D1-T179 mutants on TAP and High

Salt media.

154

3.3.3 Oxygen evolution measurements of WT and D1-T179 mutants

To determine the effect of the D1-T179 mutations on photosynthetic oxygen evolution, steady-state rates of oxygen evolution were measured using a Clark-type oxygen electrode at saturating light intensities (850 µmol photons m-2 s-1 of red light at

650 nm). While all the mutants had reduced levels of oxygen evolution activity, concomitant with the growth phenotype, the biggest decreases were observed in the cases of the D1-T179N and I substitutions. The D1-T179S, D, H, N and I mutants had 71%,

64%, 60%, 43% and 37% of the oxygen evolution activity of the wild-type control, respectively (Table 3.3).

155

-1 -1 Strain Oxygen evolved (µmol O2 mg Chl hr ) Percentage of WT

WT 107 ± 5 100

D1-T179S 76 ± 1 71

D1-T179D 68 ± 1 64

D1-T179H 64 ± 2 60

D1-T179N 46 ± 2 43

D1-T179I 40 ± 3 37

Table 3.3 Rates of steady-state oxygen evolution measured in whole cells of the complemented WT and D1-T179 mutants.

156

3.3.4 Basal Chl fluorescence (F0) and flash-induced Chl fluorescence relaxation

kinetics

Basal Chl fluorescence (F0) levels measured at low light intensities before administering the saturating flash, were higher in the D1-T179N and I mutants compared to the WT (Fig. 3.3). This was reflective of either a larger number of inactive PS II centers or less efficient excitation energy transfer between the antenna complexes and the

PS II reaction center. The Chl fluorescence yield obtained from whole cells of WT

Chlamydomonas after a saturating flash increases rapidly due to the reduction of QA following primary charge separation and subsequently relaxes due to the reoxidation of

- QA in the dark (Vass et al., 1999) (Fig. 3.3). In Synechocystis 6803 cells this relaxation is

- characterized by three main lifetime components. In this study, the reoxidation of QA in

WT Chlamydomonas (which may be compared to the D1-Q130E cyanobacterial mutant) and mutants has been described in much the same way, using a fast, middle and slow lifetime component. The fast and middle components contribute to ~75% and ~15% of

- the total decay and are characteristic of forward electron transfer from QA to QB electron transfer when plastoquinone is either present or absent from the QB site at the time of the flash (Table 3.4). The slow component is representative of the back reaction or charge

- recombination of the S2 state of the water-oxidizing complex with QA , which in turn

- - depends on the equilibrium between QA QB and QAQB . Although mutations of the D1-

T179 residue are expected to primarily affect the properties of ChlD1, which has been implicated as being the primary electron donor in PS II, the slightly slower lifetimes associated with the fast phase and middle components in the D1-T179 mutants suggest

157 that the amino acid substitutions might also impact the stability of PS II structure and QA to QB ET (Table 3.4). The slow component, that has a lifetime of ~3.1 s in the WT and

- represents the kinetics of charge recombination between the S2 state of the OEC and QB , was accelerated to 1.3, 1.0 and 0.9 s in the D1-T179D, N and H mutants respectively

(Table 3.4). The D1-T179S transgenic also had a slightly faster lifetime but was the most

- similar to WT as expected. Finally, the S2QB back reaction in the D1-T179I mutant was slower than the WT.

158

Fig. 3.3 Chl fluorescence decay curves measured in the absence of 20 µM DCMU.

159

Strain Fast phase Middle phase Slow phase

T1 (ms)/Amp(%) T2 (ms)/Amp(%) T3 (s)/Amp(%)

D1-T179I 0.37 ± 0.04/63 ± 1.0 21.4 ± 5.5/22 ± 0.8 5.6 ± 2.8/15 ± 1.3

D1-WT 0.27 ± 0.01/75 ± 0.7 55.0 ± 5.9/14 ± 0.4 3.1 ± 0.3/12 ± 0.5

D1-T179S 0.33 ± 0.02/71 ± 0.9 26.5 ± 2.5/16 ± 0.5 2.2 ± 0.5/13 ± 0.5

D1-T179N 0.29 ± 0.01/71 ± 0.5 18.0 ± 4.5/17 ± 0.7 1.3 ± 0.2/12 ± 0.9

D1-T179D 0.36 ± 0.01/74 ± 0.8 36.1 ± 3.3/16 ± 0.8 1.0 ± 0.1/11 ± 0.4

D1-T179H 0.32 ± 0.06/75 ± 0.9 22.9 ± 10.6/15 ± 0.9 0.9 ± 0.3/10 ± 1.1

Table 3.4 Chl fluorescence decay kinetics in the absence of 20 µM of DCMU.

160

3.3.5 Chl fluorescence decay in the presence of QB site inhibitors

Modifying the free energy level of the donor/acceptor pair impacts the charge recombination process in PS II and in turn influences the kinetics of flash-induced Chl fluorescence decay. More specifically, changes in the midpoint potential of the primary

- electron donor or acceptor induce changes in QA reoxidation kinetics. In the presence of

QB site inhibitors such as DCMU, ET from QA to QB is inhibited and the reoxidation of

- QA ensues via back reactions with donor side components such as the S2 state of the

- OEC (S2QA recombination). An acceleration of the Chl fluorescence decay kinetics in the presence of DCMU was observed in all the mutants in which the substituted amino acid was more polar than threonine (WT), regardless of the charge of the amino acid side

- chain (Radzicka and Wolfenden, 1988) (Fig. 3.4). The back reaction from QA to S2 was accelerated from 0.25 s in the wild type (D1-T179) to 0.22, 0.12, 0.09 and 0.07 s in the

D1-T179S, N, D and H mutants respectively (Table 3.5). In these mutants, the free

+ - energy level of the P680 PheoD1 state is presumably shifted to more positive values which leads to faster overall charge recombination rates than in the WT. Conversely, when the

+ - free energy of the P680 PheoD1 state is shifted to more negative values, as in the mutant

- with the non-polar substitution, D1-T179I, the rate of S2QA charge recombination is retarded to 0.36 s. These findings are in agreement with the previous data obtained for the

+ - D1-E130L mutant of Chlamydomonas where the free energy of the P680 PheoD1 state is shifted to more negative values and charge recombination is slower than in WT

(Rappaport et al., 2005).

161

Fig. 3.4 Chl fluorescence decay curves measured in the presence of 20 µM DCMU. The curves shown were normalized to the same initial amplitude.

162

Strain Fast phase Slow phase

T1 (ms)/Amp(%) T2 (s)/Amp(%)

D1-T179I 6.8 ± 2.5/12.0 ± 0.8 0.36 ± 0.02/88.0 ± 0.8

D1-WT 8.3 ± 0.8/6.2 ± 0.4 0.25 ± 0.01/93.8 ± 0.4

D1-T179S 3.4 ± 0.7/5.7 ± 0.7 0.22 ± 0.01/94.3 ± 0.7

D1-T179N 4.5 ± 1.5/7.2 ± 1.5 0.12 ± 0.00/92.8 ± 0.7

D1-T179D 2.5 ± 0.4/5.3 ± 0.5 0.09 ± 0.00/94.7 ± 0.5

D1-T179H 1.7 ± 0.3/5.5 ± 0.9 0.07 ± 0.00/94.5 ± 0.9

Table 3.5 Chl fluorescence decay kinetics in the presence of 20 µM of DCMU.

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3.3.6 Thermoluminescence properties of WT and D1-T179 mutants

+ - Since the ET reactions of PS II are reversible, charge separated states such as P680 QA can recombine to regenerate the ground state. This recombination can occur in three possible ways that encompass a direct non-radiative (direct electron tunneling) pathway

+ - and an indirect pathway in which the P680 PheoD1 state is formed as an intermediate

(Rappaport et al., 2005; Cser K and Vass I., 2007). This radical pair can decay in a non-

3 * radiative manner via direct recombination (or through the formation of P680 ), or in a

1 * radiative fashion giving rise to the P680 Phe state, which decays via light emission and is the origin of the phenomenon of photosynthetic thermoluminescence (TL). The TL band generated from WT Chlamydomonas cells that arises due to charge recombination

- between the QB and S2 radical pair has a maximum emission at 28 C and is called the B band (Inoue, 1996). Previous studies on Chlamydomonas and Synechocystis mutants in

+ - which the free energy of the P680 PheoD1 primary radical pair was altered by site-directed mutagenesis of amino acid residues usually in close proximity of the cofactors, have demonstrated that the intensity and peak temperatures of the B band (or the Q band that appears instead of the B band in the presence of DCMU) are extremely sensitive to

+ - changes in the free energy of the P680 PheoD1 primary radical pair even when the overall

* - free energy gap between P680 and the stable charge separated state S2QB , is unchanged

(Rappaport et al., 2005; Cser and Vass, 2007). The TL curves for the WT and mutants obtained in the absence of ET inhibitors are shown in Fig. 3.5. Significantly reduced B- band intensities and lower peak temperatures were observed in the D1-T179D, H and N mutants that had accelerated charge recombination rates and was presumably reflective of

164

+ - a change in free energy level of the P680 PheoD1 pair (Table 3.7). The peak temperature of the B band was shifted from ~28 C in the WT to ~21, 20, and 19 C in the D1-

T179D, H and N mutants respectively. The D1-T179I mutant that had the slowest rate of charge recombination as measured via Chl fluorescence decay kinetics exhibited a TL

- band at ~16 ºC and another band at ~40 ºC (S2/3QB recombination). The band at ~16 ºC

- - most likely represents charge recombination between S2QA rather than S2QB . In the

- absence of ET inhibitors, S2QB charge recombination should be more prevalent than

- S2QA recombination as mentioned above. However inefficient excitation energy transfer and decreased primary charge separation yields most probably result in an increase of non-QB-reducing PS II centers in the D1-T179I mutant, causing the appearance of a TL

- band corresponding to S2QA recombination even in the absence of ET inhibitors. The raw intensity of the TL band was also reduced, to ~10% of the WT, due to the low yields of charge separation in this mutant as is evident from decrease oxygen evolution rates and variable fluorescence.

165

Fig. 3.5 Thermoluminescence characteristics of the WT and D1-T179 mutants measured in the absence of 20 µM DCMU.

166

The TL bands obtained for the WT and mutants in the presence of DCMU when QA to QB ET is inhibited are shown in Fig. 3.6. Under these conditions, the dominant TL

- peak, designated the Q band, arises due to S2QA recombination and consistently appears at 5-6 ºC in the WT in the presence of 0.5 µM nigericin (Inoue, 1996). Analogous to the modifications seen in the B band, the intensity of the Q band was also reduced in the D1-

T179D, H and N mutants which had highly polar residues present in position D1-179 when compared to the WT. Additionally, the peak temperatures of the Q band in the D1-

T179D, H and N mutants were shifted to lower temperatures than in the WT appearing between 0-3 C compared to 5-6 C in the WT. The D1-T179S mutant had similar TL characteristics as the WT suggesting that the D1-T179S mutation did not significantly impact the redox potential of P680, while the D1-T179D, H and N mutations did. Lastly, the Q band of the D1-T179I mutant had a slightly higher peak temperature of 8 C when compared to the WT, although the raw TL intensity was reduced due to reduced yields of primary charge separation.

167

Fig. 3.6 Thermoluminescence characteristics of the WT and D1-T179 mutants measured in the presence of 20 µM DCMU.

168

3.4 Discussion

One of the most striking differences between type II bacterial RCs and PS II is that excitation of PS II leads to the formation of a highly oxidizing primary donor cation that can drive water oxidation. At the level of primary protein sequence of the BRC and PS II

RC polypeptides, no substantial differences are seen in acceptor side residues surrounding the RC Pheos. However, there are obvious differences between the two RC types in the residues located near the donor side cofactors, especially in the vicinity of the monomeric Chls. For example, the orientation and hydrogen bonding properties of the

BRC residues located between L-E104 and L-W100 which are adjacent to the BPheoL are largely conserved in the corresponding PS II residues, D1-E130 and D1-Y126. However, while histidine residues ligate the BChl monomers in the BRC, the corresponding PS II residues are non-ligating amino acids. The residues closest to the monomeric Chls ChlD1 and ChlD2 in PS II are D1-T179 and D2-I178 respectively. Moreover, a comparison of all available D1 protein sequences show that threonine (and not histidine as in the progenitor) is completely conserved at position 179. Previously, studies involving the substitution of D1-T179 with histidine and glutamine residues in cyanobacteria were shown to modify the spectroscopic properties of ChlD1 (Schlodder et al., 2008a,b).

However, the role of D1-T179 in determining the redox properties of the primary electron donor was not addressed. Through comparisons of flash induced (3P-1P) absorbance difference spectra of the mutants and WT, it was ascertained that the RC triplet state that is generated upon charge recombination is localized on the monomeric ChlD1 at cryogenic temperature. In addition, the lowest exciton transition was shown as being localized on

169

ChlD1 and not on the Chl special pair as in the bacterial RC or PS I. These results provided more evidence that ChlD1 is the primary electron donor in PS II.

In this work, the effects of replacing this highly conserved threonine with residues that retain either all, some or none of the neutral and polar properties of threonine, are studied in detail. The resulting transgenics were examined for their capacity to perform photoautotrophic growth and evolve oxygen. Additionally, the effects of the mutations on

+ - the redox properties of P680 and on the free energy level of the P680 PheoD1 radical pair were studied through flash-induced Chl fluorescence and thermoluminescence measurements in the absence and presence of ET inhibitors.

The ability of all the D1-T179 mutants to carry out photoautotrophic growth on high salt media was affected to varying extents with the most severe growth phenotypes being seen in the D1-T179N and D1-T179I mutants (Fig. 3.2). As expected, the steady-state oxygen evolution rates measured in whole cells were also affected in all the mutants, with the D1-T179N and D1-T179I mutants having only ~43% and ~37% of the WT activity respectively (Table 3.2). The levels of basal Chl fluorescence (F0), variable fluorescence

(Fv = Ft - F0) and maximum fluorescence (Fm) were obtained from the Chl fluorescence decay curves for the WT and mutants (Fig. 3.3). The ratio of Fv/Fm is reflective of the quantum efficiency of PS II and was found to be 0.66 in the WT. The D1-T179S, D, H, N and I transgenics had substantially reduced levels of variable fluorescence and Fv/Fm values of 0.57, 0.50, 0.51, 0.38 and 0.29 respectively, indicative of reduced PS II efficiency, corroborating the decreased oxygen evolution rates and photosynthetic growth seen in the mutants. Furthermore, the D1-T179N and D1-T179I mutants had elevated

170 basal Chl fluorescence (F0) and reduced variable Chl fluorescence levels compared to the

WT. The value of F0 is reflective of the Chl fluorescence emanating from the PS II antenna in competition with excitation energy transfer to the RC. Elevated F0 levels are indicative of inefficient excitation energy transfer from the proximal PS II antenna complex to the RC core. We calculated the predicted coordination distances for the mutants by way of in silico mutagenesis of the D1-T179 residue using the PyMOL

(Delano Scientific) software and observed a correlation between increased predicted amino acid residue proximity to ChlD1 and F0 level. This suggested that the efficiency of excitation energy transfer to the RC Chls from the proximal PS II antenna was compromised with decreasing predicted coordination distance as in the D1-T179I and N mutant PS II RCs (Table 3.6). However, elevated F0 levels were not observed for the D1-

T179D mutant which also had an increased predicted proximity to ChlD1, possibly because of complimentary structural rearrangements.

171

D1-179 Possible Coordination Distance (Å)

Ile 3.46 Thr (WT) 4.95 Ser 4.87 Asp 3.69 His 5.24 Asn 3.71

Table 3.6 Possible coordination distances predicted through backbone-dependent in silico mutagenesis of the D1-T179 residue. This was based on the PDB coordinates 3BZ1, of the PS II crystal structure described in Guskov et al., 2009.

172

The effects of replacing the residue at position 179 of the D1 protein (with residues either more or less polar than in the wild-type) on the redox properties of the monomeric

ChlD1 of the active branch were studied using flash-induced Chl fluorescence and thermoluminescence measurements. While the former collectively reflects radiative and non-radiative charge recombination pathways, luminescence in the latter originates from the indirect radiative pathway. Both methods however, are sensitive to changes in the free energy level of PS II redox components and can be used to study the effects of mutations that alter the redox properties of the core RC pigments. The dominant pathway

+ - for charge recombination in PS II is through the P680 PheoD1 primary radical pair

+ - (Rappaport et al., 2005; Cser et al., 2008). Charge recombination between P680 PheoD1 is further classified under radiative and non-radiative pathways which originate via the re-

1 + - formation of the P680* state from [P680 PheoD1 ] and from the direct recombination of the

3 + - 1 + - [P680 PheoD1 ] and [P680 PheoD1 ] radical pairs, respectively. In addition, the direct

+ - recombination of P680 with QA can occur in a non-radiative manner. The yield of the different charge recombination pathways of PS II is modulated by the free energy gaps between the PS II ET components.

The relative intensity of the TL band is related to the free energy level of the primary radical pair which in turn determines the preferred charge recombination pathway and the

* + propensity of re-forming P680 . Shifting the mid-point potential of either the P680 /P680 or

- PheoD1/PheoD1 couple to more negative values decreases the free energy gap between the

* * primary radical pair and the P680 , thereby increasing the probability of reforming P680 and the TL yield. Conversely, if the mid-point potential of the primary redox couples are

173

+ * shifted to more positive values, then the free energy gap between P680 PheoD1 and P680

* increases, thereby decreasing the possibility of re-forming P680 upon charge recombination. This leads to a consequent decrease in TL yield and an increase in non- radiative recombination to the ground state or re-separation to the first stable charge

+ - separated state P680 QA .

The site-directed mutagenesis of the D1-T179 residue has a marked effect on TL intensity as seen in Figs. 3.5 and 3.6. Given that the location of this residue is close to the

ChlD1 on the donor side, the mutational effects on the TL bands are due most likely to altered energetics of the primary electron donor. Furthermore, the overall free energy gap

- - between S2 and QA (QB ) is not expected to be altered by the mutations. The modulation of the normalized TL intensity in the D1-T179 mutants despite no change in the energetic

- - gap between S2 and QA (QB ) confirms that the free energy difference between P680* and

+ - P680 PheoD1 is an important determinant of TL emission (intensity) in agreement with

+ - previous results. Hence, changes in the energetic gap ΔG(P680*↔P680 PheoD1 ), induced by the D1-T179 mutations can be inferred from the TL intensities of the respective mutants relative to the wild-type. However, TL intensity is also dependent on the yield and efficiency of primary charge separation and on the number of functional PS II centers. In order to negate the contribution of these effects, the raw TL intensities (area under the TL curve) obtained for each strain were normalized to the initial amplitudes

(Fv/F0) of the Chl fluorescence transients measured in the presence of DCMU, in the

- same strain (Table 3.7). The changes in the free energy differences of S2QB recombination in the D1-T179 mutants relative to the WT were then calculated using the

174 normalized intensity of the TL bands as Δ(ΔG) = kT.ln(TLmutant/TLWT) as performed in

Cser and Vass, 2007. This calculation yielded an overall decrease in the free energy for

- * + - S2QB recombination and a consequent increase in ΔG(P680 ↔P680 PheoD1 ) of about 16 to

25 meV in the D1-T179D, N and H mutants compared to the WT (Table 3.7 and Fig.

- 3.7). Moreover, this was accompanied by shifts in the peak positions of the B band (S2QB recombination) to lower temperatures compared to the WT, consistent with previous findings that the peak temperature of thermoluminescence is determined by the free

+ - - - energy gap between P680 PheoD1 and S2QB (or S2QA in the presence of QB site inhibitors). The normalized B band intensity of the D1-T179S mutant showed little variation from the WT suggesting no change in the free energy level of the primary radical pair in this mutant.

175

B band (S Q -) 2 B Normalized TL ΔG(P +Pheo -) Strain Peak Intensity 680 D1 temperature Relative to WT (meV) (C) WT 28 ± 0.2 1

D1-T179S 25 ± 1.8 1.06 ± 0.01

D1-T179D 21 ± 0.9 0.53 ± 0.02 -16.3

D1-T179H 20 ± 0.6 0.38 ± 0.02 -24.8

D1-T179N 19 ± 0.8 0.40 ± 0.04 -23.5

Table 3.7 Free energy changes related to charge recombination in the D1-T179 mutants.

176

While thermoluminescence is a result of indirect radiative charge recombination, flash-induced Chl fluorescence is the result of the sum of both radiative and non-radiative processes and can be conveniently measured by following the relaxation kinetics of flash- induced Chl fluorescence in the presence of DCMU. The kinetics of this process were

Chl fluorescence decay kinetics were accelerated in the D1-T179D, H and N mutants

+ - consistent with an increase in the free energy level of P680 PheoD1 (Fig. 3.4, Table 3.4).

- S2QA charge recombination in the D1-T179I mutant, on the other hand, was slightly retarded from 0.25 s in the WT to 0.36 s in the mutant, suggesting a decrease in the free

+ - energy level of P680 PheoD1 compared to WT. A straightforward explanation of these

- + - data is that the free energy difference between S2QA and P680 PheoD1 in the D1-T179 mutants is modified compared to the WT. The change in the overall free energy of the

- + S2QA recombination due to changes in P680/P680 redox potential in the D1-T179 mutants, relative to the WT, can be estimated by comparing the lifetime components of the slow phases of the mutants and WT in the presence of DCMU using the equation ΔGmutant –

ΔGWT = kT.ln(τslow, mutant/τslow, WT) and the assumption that the mutations do not affect the redox potential of QA (Vass et al., 1999). This calculation yields an increase in the overall

- free energy for S2QA recombination for the D1-T179I mutant by 9.3 meV and a decrease for the D1-T179D, H, N mutants by ~20–30 meV. In the absence of ET inhibitors, the

- slow phase of fluorescence decay arises from the recombination of the S2 state with QB ,

- - via the QA QB ↔ QAQB equilibrium. Therefore, the changing lifetime component of the slow phase, which increases in D1-T179I and decreases in the D1-T179D, H and N mutants relative to the WT can be explained by a changing free energy gap between

177

+ - - - P680 PheoD1 and S2QB in the same way as described above for S2QA charge recombination.

Hence, site-specific mutations of D1-T179 to more polar residues such as aspartate,

+ - asparagine and histidine, cause changes in the free energy level of the P680 PheoD1 primary radical pair by +16-25 meV relative to the WT as estimated by TL measurements

(Fig. 3.7). Due to the low TL yield and inefficiency of QB reduction in the D1-T179I mutant, similar calculations for the mutational changes in the free energy level of the primary radical pair could not be ascertained through TL measurements. However, since

+ - changes in the free energy level of P680 PheoD1 due to the modification of P680, also

- influences the kinetics of S2QA recombination, the decay of flash-induced Chl fluorescence measured for the D1-T179I mutant in the presence of DCMU was slower

- compared to the WT. This suggested an overall increase in the free energy for S2QA charge recombination. Similar analyses of the D1-T179D, H and N, revealed faster decay

- kinetics in the presence of DCMU suggesting a decrease in the free energy for S2QA charge recombination.

The combined results of the thermoluminescence and flash-induced Chl fluorescence decay measurements suggest that the mutagenesis of the D1-T179 residue to amino acids

+ more polar than threonine, cause a shift in the Em of P680/P680 to more positive values such that the probability of indirect radiative charge recombination decreases due to an

* + - increase in the energetic gap between P680 and the primary radical pair P680 PheoD1 .

- However the overall rate for S2QA charge recombination increases due to a decrease in

- the energetic gap between the primary radical pair and the S2QA state (Fig. 3.7). The

178 effects of the mutagenesis of D1-T179, on the redox properties of P680 provide more support to the suggestion that ChlD1 is the primary electron donor in PS II. They also suggest that the properties of the residue at this position are of paramount importance in determining the efficiencies of forward ET and charge recombination.

In conclusion, for the optimum functioning of oxygenic photosynthesis in PS II, forward ET and charge recombination must be balanced in favor of forward ET. Changes

+ in the redox properties of the P680/P680 couple to more positive values such as in the D1-

T179D, H and N mutants accelerate charge recombination and reduce the efficacy of forward ET in PS II. Therefore in PS II, in contrast to the type II BRC, having a histidine residue in closest proximity to the ChlD1 monomer of the active branch reduces the efficiency of forward ET and oxygenic photosynthesis. It is foreseeable that, while the PS

II RC was derived from and maintains majority of the properties of the BRC, the residue near the monomeric ChlD1 was substituted (by threonine) during the evolution of oxygenic photosynthesis, to provide for an efficient balance between the ability to generate a highly oxidizing primary donor cation for water oxidation, and forward and back electron transfer.

179

Fig. 3.7 Scheme of charge recombination pathways in PS II. The double headed blue arrows show the direction of free energy changes induced by the mutations of the D1-

T179 residue.

180

Chapter 4

Site-Directed Mutagenesis of D2-Q129 Alters the Redox Potential of QB and Charge

Recombination in Photosystem II

4.1 Introduction

Photosystem II (PS II) is a multi-subunit pigment-protein complex found in oxygenic photoautotrophs that catalyzes the light-driven oxidation of water and reduction of plastoquinone. The structure of PS II from thermophilic cyanobacteria is now available at

2.9 Å resolution, along with several lower resolution crystal structures (3-3.8 Å) (Zouni et al., 2001; Ferreira et al., 2004; Loll et al., 2005; Guskov et al., 2009). PS II has two branches of redox active cofactors, only one of which is active in electron transfer. It is well established that the primary electron acceptor in PS II is a pheophytin molecule,

PheoD1, which resides on the active branch (Klimov et al., 1977; Klimov and Kravnoski,

1981; Klimov, 2003; Groot et al., 2005; Holzwarth et al., 2005). Similar to the purple bacterial reaction center (BRC), there are two symmetry-related pheophytins (Pheo) in PS

II (Fig. 4.1). Prior to the availability of a high resolution PS II crystal structure, the presence of a hydrogen bonding interaction between the chloroplastic D1-E130 residue and the ring V carbonyl group of PheoD1 was established using a

181 combination of site-directed mutagenesis and EPR spectroscopy of the Pheo anion radical

(Dorlet et al., 2001). Mutations that were designed to weaken or remove this hydrogen bond affected the g tensor of the Pheo anion radical. An upward shift of the gx component was seen in the D1-E130Q, H and L Chlamydomonas mutants that had decreasing hydrogen bonding capacities compared to wild-type. The greatest shift in gx value was observed for the D1-E130L mutant, in which this hydrogen bond interaction was abolished. Subsequently, Chl fluorescence decay kinetic data obtained for the D1-E130L

Chlamydomonas and D1-Q130L cyanobacterial mutants showed that the loss of the

- hydrogen bond interaction to PheoD1 resulted in a longer lifetime for S2QA charge recombination (Cuni et al., 2004; Rappaport et al., 2005; Cser and Vass, 2007).

Additionally, thermoluminescence measurements on the same mutants revealed that the

+ - + - energetic gap between the primary radical pair P680 PheoD1 and P680 QA increased as a result of a leucine substitution and the loss of a hydrogen bond donor to PheoD1

(Rappaport et al., 2005; Cser and Vass, 2007). It was inferred that mutagenesis of D1-130 to amino acids that weakened the hydrogen bonding interaction to PheoD1 shifted the

- midpoint potential of the PheoD1/PheoD1 couple to more negative values, making forward

ET transfer from PheoD1 to QA less probable. Hence, the D1-E130L PS II mutation effectively results in an impairment of forward ET and a reduction in primary charge separation yield.

As mentioned previously, PheoD2 is related to PheoD1 by C2-symmetry but does not take part in primary ET in PS II. The role of PheoD2 and its protein environment is not as well understood as the role and protein environment of PheoD1. There is some evidence to

182 show that PheoD2 may be excitonically coupled to the central multimeric RC chlorins and hence be involved in excitation energy equilibration within the RC complex (Durrant et al., 1995). The substitution of PheoD2 with a chlorophyll (Chl) molecule has been shown to result in substantial impairment of PS II (Xiong et al., 2004). Pigment substitution was carried out via the site-specific mutagenesis of D1-L210 to histidine which enabled the

2+ coordination of a Mg atom in the center of the PheoD2 macrocycle ring converting it to a Chl. Interestingly, the perturbation of this inactive branch cofactor as in the D1-L210H mutant drastically reduced oxygen evolution rates and primary charge separation yields.

Moreover, the circular dichroism (CD) spectrum of D1-L210H RCs had obvious changes in the red region of the spectrum at 669 and 682 nm when compared to wild-type. These changes were interpreted as resulting from the combined loss of excitonic interactions between PheoD2 and the RC multimer pigments, and the addition of new excitonic interactions between the substituted Chl and the neighboring pigments. CD spectra of

1 pigment substituted PS II RCs, in which PheoD2 was replaced by modified Pheo (13 -OH-

Pheo), showed changes in similar regions of the CD spectrum as the D1-L210H mutant

(Germano et al., 2001). These observations were consistent with involvement of PheoD2 in excitonic interactions with other RC chlorins. The D1-L210H mutant also showed increased levels of low temperature Chl fluorescence emission consistent with the possible redistribution of the excited-state equilibrium among the core pigments of the PS

II RC (Xiong et al., 2004). Hence, the possible function of PheoD2 may be inferred from the results of the D1-L210H mutant as being necessary to ensure the proper distribution

183 of excitation energy among pigments of the multimer complex such that efficient active branch charge separation can take place.

As earlier described, the protein environment of PheoD1 is of paramount importance in determining the yield of primary charge separation in PS II. The D1-130 residue of the cyanobacterial PS II is 3 Å away from PheoD1 and therefore can engage in a hydrogen bonding interaction with the PheoD1 head group. The inactive branch residue analogous to

D1-E130 in chloroplastic PS II is D2-Q129. D2-Q129 has also been suggested as being a possible hydrogen bond donor to the head group of PheoD2 (Xiong et al., 1996; Kern and

Renger, 2007). However, analysis of the most recent PS II crystal structure from the cyanobacterium Thermosynechococcus elongatus at 2.9 Å resolution reveals that the side chain of D2-Q129 is 3.9 Å away from the PheoD2 head group and therefore may not be in close enough proximity to serve as a viable hydrogen bond donor (Fig. 4.1). In the RCs of purple bacteria, L-E104 (analogous to D1-E130 in chloroplastic PS II) is highly conserved and is known to hydrogen bond to BPheoL (Bylina et al., 1988; Deisenhofer and Michel, 1989). The corresponding inactive (M) branch residue shows little conservation and M, T, V, A and I residues have been known to occur. In contrast, a comparison of all 152 protein sequences available for the D2 polypeptide of PS II from a variety of oxygenic photoautotrophs shows that D2-Q129 is completely conserved across all sequences.

In general, little is known about the specific roles of acceptor-side inactive branch residues in PS II. In this study, we characterized the role of the highly conserved acceptor side residue D2-Q129. In contrast to its well-studied active branch counterpart (D1-

184

E130), no specific function had previously been assigned to D2-Q129 based on experimental data. We applied a mutagenesis approach in which D2-Q129 was replaced by a non-conservative hydrophobic leucine residue (D2-Q129L) or a conservative histidine residue (D2-Q129H) in Chlamydomonas reinhardtii. For comparison, we also generated the analogous D1-E130L single mutant as well as the D1-E130L/D2-Q129L double mutant. The resulting transgenics were analyzed by their photoautotrophic growth characteristics, oxygen evolution rates, Chl fluorescence induction and decay kinetics, thermoluminescence properties and sensitivity to photoinhibitory light treatment. Our Chl fluorescence decay analysis and thermoluminescence data both showed that the mutagenesis of D2-Q129 to either conservative or non-conservative amino acid residues,

- - led to faster rates of S2QB charge recombination while no change was apparent in S2QA charge recombination. Furthermore, mutagenesis of D2-Q129 also caused increased susceptibility to photoinhibitory light treatment and photooxidative damage indicating that inactive branch residues may help in optimizing PS II function under a variety of growth conditions.

Our results also demonstrate that the amino acid residue in position 129 of the D2 protein has a major influence on the redox potential of QB. This result was unanticipated given that the analogous active branch residue, D1-E130, influences the redox properties of PheoD1. The disparate effects of the analogous D1-E130L and D2-Q129L mutations shed light on the extent of asymmetry between the two cofactor branches in PS II.

Moreover, this study documents the first finding of a single amino acid substitution of the

D2 protein which alters the redox potential of QB. Previous mutations that have been

185 shown to alter QB redox potential have most often been attributed to the D1 protein

(Ohad and Hirschberg, 1992; Etienne and Kirilovsky, 1993; Nixon et al., 1995; Constant et al., 1996; Lardans et al., 1998, Minagawa et al., 1998; Sane et al., 2002; Rose et al.,

2008).

Given that the D2-Q129 residue is located close to PheoD2, we wanted to unequivocally determine whether or not any of the inactive branch cofactors were

- involved in the charge recombination of QB with S2, in the D2-Q129 mutants. For this, we generated the D1-E130L/D2-Q129L double transgenic strain (or LL mutant) with concurrent mutations in the amino acid residues associated with both the active and inactive PS II RC cofactor branches. Our data show that the D1-E130L/D2-Q129L double mutant manifests the combined effects of the two single mutations. This indicated

- that the acceleration of charge recombination from QB to the S2 state of the water oxidation complex, induced by the D2-Q129 mutation, in fact involves the active branch and not the inactive branch cofactors. To our knowledge, this is the first report of simultaneously combining site-directed mutations of both the D1 and D2 polypeptides and of the ―active‖ and ―inactive‖ branches of the PS II RC complex in one transgenic strain.

186

Fig. 4.1 Structure of the acceptor side of Photosystem II. Inter-atomic distances are indicated by dashed black lines. The amino residues forming the QA and QB binding pockets are shown as green and cyan sticks respectively. Amino acid residues of the D1- de helix that are known to play a role in determining the redox potential of QB, are shown as orange sticks and the corresponding residues of the QA site are shown in pale brown.

This view of PS II is in an orientation parallel to the plane of the membrane and was created using PyMOL (2006 DeLano Scientific LLC) and the 3BZ1 PS II crystal structure.

187

4.2 Materials and Methods

4.2.1 DNA constructs

Site-directed mutations were introduced into the Chlamydomonas reinhardtii psbA

(encodes the D1 PS II RC protein) and psbD (encodes the D2 PS II RC protein) genes using the Quik-Change Site-Directed mutagenesis kit from Stratagene. The pBA155 and pBD202 (a gift of Dr. Jun Minagawa) vectors containing the psbA and psbD genes were used as the templates for site-directed mutagenesis. Both plasmids also contained the aadA gene that confers resistance to the antibiotics spectinomycin and streptomycin

(Goldschmidt-Clermont, 1991). A mutation was introduced at position 130 of the D1 protein in which the glutamate codon GAG was replaced with a leucine codon CTT using the D1-E130-F and R primers shown in Table 4.1. Similarly, two different point mutations were introduced at position 129 of the D2 protein. The glutamine-129 codon

CAG was mutagenized either to CTG (leucine substitution) or CAC (histidine substitution) using primer sets D2-Q129L-F and R and D2-Q129H-F and R respectively

(Table 4.1). The psbA and psbD mutations were confirmed by PCR and sequencing of the mutagenized plasmids using the D1-5‘/3‘UTR and D2-5‘/3‘UTR primers sets respectively (Table 4.1).

188

Primer Sequence (5'3')

D1-E130L-F CTACATGGGTCGACTTTGGGAATTATC

D1-E130L-R GATAATTCCCAAAGTCGACCCATGTAG

D2-Q129L-F GGTTTCATGCTTCGTCTGTTTGAAATTGCTCGTTC

D2-Q129L-R GAACGAGCAATTTCAAACAGACGAAGCATGAAACC

D2-Q129H-F TTGGTTTCATGCTTCGTCACTTTGAAATTGCTCGTTCAG

D2-Q129H-R CTGAACGAGCAATTTCAAAGTGACGAAGCATGAAACCAA

D1-5'UTR GGACGTAGGTACATAAATGTGCTAGGTAAC

D1-3'UTR CCTGCCAACTGCCTATGGTAGCTATTAAGT

D2-5'UTR GTGATGACTATGCACAAAGCAGTTCTAGTCCC

D2-3'UTR CAAGCACTCATGTGATTTTTAGCCCCAAAGGG

Table 4.1 List of primers used for site-directed mutagenesis of amino acid residues D1-

E130 and D2-Q129.

189

4.2.2. C. reinhardtii chloroplast transformation

The D1-E130L plasmid was introduced into the psbA deletion strain CC-4147

(Chlamydomonas Genetics Center, Duke University) by particle gun bombardment to generate the D1-E130L Chlamydomonas mutant. The pBD202, D2-Q129L and D2-

Q129H vectors were introduced into a psbD deletion mutant, ΔD2-2-6 (a gift of Dr. Jun

Minagawa) by particle bombardment to generate the complemented wild type (WT) and

D2-Q129L and D2-Q129H mutant strains. In addition, the D1-E130L and D2-Q129L plasmids were simultaneously introduced into a {ΔpsbA, ΔpsbD} double deletion strain

(a gift from Dr. Yves Choquet, Minai et al., 2006) to generate the

D1-E130L/D2-Q129L (LL) double mutant with substitutions at positions 130 and 129 of the D1 and D2 proteins respectively, to leucine.

Following bombardment, the cells were plated on Tris-acetate-phosphate (TAP) medium (Harris, 1989) containing 100 µg/mL spectinomycin and incubated at 22 C under dim light. Putative transformants were selected on the basis of spectinomycin and streptomycin resistance as described in Chapter 3. DNA extracted from the transgenics via the Chelex-100 extraction method (Cao et al., 2009) was used as template for DNA sequence confirmation of the mutagenized psbA and psbD genes using PCR primers D1-

5‘/3‘UTR and D2-5‘/3‘UTR shown in Table 4.1.

190

4.2.3 Growth of Chlamydomonas WT and mutant (D1-E130L, D2-Q129L, D2-

Q129H and D1-E130L/D2-Q129L) cells

All strains were maintained on TAP agar plates containing 100 µg/mL spectinomycin and 50 µg/mL ampicillin and were grown up to mid to late log phase for subsequent experiments as described in Chapter 3. The complemented wild-type (WT) and mutant strains were normalized on the basis of cell number and assayed for photoautotrophic growth on high salt media under low light conditions (~30 µmol light m-2 s-1) as described in Chapter 3. In addition, the WT, D2-Q129L and D2-Q129H mutants were also grown on solid photoautotrophic media under high light conditions (~500 µmol light m-2 s-1).

4.2.4 Oxygen evolution measurements

Oxygen evolution activity of wild-type and mutant cells was measured under saturating conditions of ~850 μMol light m-2 s-1 of 650 nm light, using a Clark-type oxygen electrode (Hansatech Instruments) as described in Chapter 3. The Chl concentration used was ~10 µg Chl/mL.

4.2.5 Flash-induced Chl fluorescence induction and decay kinetics

Chl a fluorescence induction transients of wild-type and mutant cells were measured with a pulse-modulated fluorometer (FL 3500, Photon Systems Instruments). Before the measurements the cells were resuspended in TAP medium at a Chl concentration of 5

µg/mL and dark adapted for 10 min.

191

The decay of Chl a fluorescence induced by a single saturating flash, due to the

- reoxidation of QA , was assayed in whole cells in the presence and absence of 20 μM 3-

(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) using a pulse modulated fluorometer

(Photon Systems Instruments) in the 100 μs–100 s time range. The samples were prepared as mentioned above. Multicomponent deconvolution of the fluorescence decay curves in the absence of DCMU was done by using a fitting function with three components, fast and intermediate exponential and a slow hyperbolic component as described in Vass et al., 1999. When measured in the presence of DCMU, the fitting

- function included only two components, fast and slow phase representing QA

• recombination with TyrZ and S2 , respectively.

4.2.6 Thermoluminescence measurements

Thermoluminescence (TL) from whole cells was measured in the presence and absence of 20 μM DCMU using a thermoluminescence instrument manufactured by

Photon Systems Instruments. A cell suspension of 50 µg Chl per 50 µL in 20 mM

HEPES at pH 7.5 and 0.5 µM nigericin was spotted on a ½ inch disc prepared from

Whatman filter paper 1. The sample was dark-adapted for 2 min and then cooled to 0 ºC or -10 ºC before TL was measured in the absence and presence of DCMU respectively.

After 2 min, a single saturating flash was fired and TL was recorded upon heating the sample at a rate of 0.5 ºC/s. For p-benzoquinone (PBQ) treatment, cells were incubated with 100 µM PBQ for 10 min in the dark, then spun down and washed to remove the residual PBQ before being used for the measurement.

192

4.2.7 Photoinhibition measurements

C. reinhardtii cultures were grown as described above and resuspended in buffer A containing 0.35 M Sucrose, 20 mM HEPES, pH 7.5 and 2 mM MgCl2 to yield a Chl concentration of 1 mg Chl/mL. Cells were then broken by sonication (Biologics, Inc,

Model 300 V/T Ultrasonic Homogenizer) two times for 10s each time (pulse mode, 50% duty cycle, output power 5) on ice. Unbroken cells were pelleted by centrifugation at

3,200 x g for 1 min and the membranes harvested by centrifugation of the supernatant obtained from the previous step at 12,000 x g for 12 min. The membranes were then resuspended in fresh buffer at > 1.0 mg Chl/mL. All steps were carried out in darkness at 4

ºC. Isolated thylakoids at a Chl concentration of ~10 µg/mL were exposed to photoinhibitory (PI) light treatment (800 μmol light m-2 s-2 650 nm at 22 ºC) and assayed for residual rates of oxygen evolution after 0, 5, 10, 15, 20, 25 and 30 minutes in the presence of 20 μM 2,5-Dimethyl-1,4-benzoquinone (DMBQ), 2 mM potassium ferricyanide and 30 mM methylamine.

4.3 Results

4.3.1 Photoautotrophic growth of WT and D2-Q129L, D2-Q129H, D1-E130L

and D1-E130L/D2-Q129L cells

As discussed previously, the D1-E130 residue interacts via a hydrogen bond with the primary electron acceptor, PheoD1, on the active branch. The analogous inactive branch residue, D2-Q129, is conserved in oxygenic photosynthesis but as revealed by the most recent crystal structure, is too far away from PheoD2 to have an analogous hydrogen

193 bonding interaction. In order to gain greater insight into the structure/function role of this residue, we carried out site-directed mutagenesis to generate the D2-Q129L non- conservative and D2-Q129H conservative mutants. Furthermore the D1-E130L single and D1-E130L/D2-Q129L (LL) double mutants were constructed to compare simultaneously the effects of mutagenizing these analogous active and inactive branch residues. To determine whether mutagenesis disrupted photosynthetic capabilities, the growth characteristics of all the mutants on photoautotrophic growth media was assayed.

Cells were diluted to equal optical densities and spotted on both TAP (control) as well as

HS plates. The growth of the D2-Q129L and D2-Q129H single mutants was comparable to the complemented wild-type (WT) in photoautotrophic media under low light intensities (Fig. 4.2). However, the D1-E130L and D1-E130L/D2-Q129L mutant strains had extremely poor photoautotrophic growth compared to WT.

194

Fig. 4.2 Growth characteristics of WT, D2-Q129L, D2-Q129H, D1-E130L and D1-

E130L/D2-Q129L (LL) cells on TAP and High Salt (photoautotrophic) media under ~30

µE m-2 s-1 light.

195

4.3.2 Oxygen evolution activity

To determine the effects of the D2-Q129 mutations on photosynthetic oxygen evolution, steady-state rates of oxygen evolution were measured at saturating light intensities using a Clark-type oxygen electrode (Hansatech) as described in Chapter 3.

The light-saturated rates of oxygen evolution in the D2-Q129L and H mutants were ~70 and 90% of the WT, respectively (Table 4.2). As expected, the oxygen evolving abilities of the D1-E130L and D1-E130L/D2-Q129L double mutants were severely reduced, being only ~20 and 10% of WT rates. In general, the extent of photoautotrophic growth in the mutants closely correlated with their oxygen evolving capacities.

196

-1 -1 Strain Oxygen evolved (µmol O2 mg Chl hr ) Percentage of WT

WT 113 ± 3.0 100

D2-Q129L 78 ± 2.6 69

D2-Q129H 101 ± 4.4 89

D1-E130L 25 ± 1.3 22

D1-E130L/D2-Q129L 10 ± 2.0 9

Table 4.2 Steady-state oxygen evolution rates of WT, D2-Q129L, D2-Q129H, D1-E130L and D1-E130L/D2-Q129L cells. The average oxygen evolution rate is based on 6-9 measurements from at least 3 independent cultures.

197

4.3.3 Flash-induced Chl fluorescence relaxation kinetics

Administration of a single-turnover saturating flash to Chlamydomonas cells leads to a transient increase in the Chl fluorescence yield due charge separation and the consequent accumulation of reduced QA. This is followed by a decay in Chl fluorescence

- due to the reoxidation of QA in the dark which can be fitted using three lifetime components and their respective amplitudes (Vass et al., 1999). Fig. 4.3 shows the flash- induced Chl fluorescence decay transient obtained in WT and mutant cells in the absence of ET inhibitors. The fast and intermediate lifetime components contribute to ~90% of

- the total decay and represent forward ET from QA to QB when plastoquinone (PQ) is either present or absent from the QB site at the time of the flash (Table 4.3). The slow component, on the other hand, contributes to ~10% of the total decay and represents

- charge recombination of the S2 state of the OEC, with QB which in turn is dependent on

- - - the equilibrium between QA QB and QAQB . The mean lifetime for forward ET from QA to QB is ~0.29 ms in the WT and was relatively unaltered in the D2-Q129L and H mutants being ~0.26 and 0.29 ms respectively. The intermediate lifetime component in the D2-Q129 mutants was also similar to the WT at ~35 ms. However, the total contribution of forward ET to the overall Chl fluorescence decay measured as the sum of the relative amplitudes of the fast and intermediate lifetime components, was significantly reduced in the D2-Q129L and H mutants to ~73 and 77% of the total respectively, relative to ~90% of the total in WT.

Conversely, the contribution of the slow component, which represents charge

- recombination between S2 and QB , showed a ~2 to 2.5-fold increase compared to WT.

198

The amplitude of the slow lifetime component is ~10% in the WT and increased to ~28 and 23% in the D2-Q129L and H mutants respectively. Furthermore, the lifetime

- component for S2QB charge recombination was accelerated ~4 to 6 fold from ~3.3 s in

WT, to 0.54 and 0.73 s in the D2-Q129L and H mutants respectively. These data indicated that while the mutagenesis of D2-Q129 did not impact the rate forward ET, it

- caused an acceleration of S2QB charge recombination. Moreover, the probability of forward ET decreased in the D2-Q129L and D2-Q129H mutants compared to WT, while

- the probability of S2QB charge recombination increased, suggesting that the overall free

- energy of S2QB charge recombination decreases as a consequence of D2-Q129 mutagenesis.

The Chl fluorescence decay kinetics obtained for the D1-E130L mutant were similar to those previously observed (Rappaport et al., 2005; Cser and Vass, 2007). The rate of forward ET was slower and its relative contribution decreased in D1-E130L compared to

- WT. The mean lifetime of forward ET from QA to QB was 0.77 ms in the mutant (0.29 ms in WT) while its amplitude decreased to 23% (72% in WT). This indicated an impairment of forward ET in the D1-E130L mutant which corroborated the poor

- photoautotrophic growth and oxygen evolution rates. Similarly, the rate of S2QB charge recombination was also substantially slower in the mutant with a lifetime of ~100 s compared to only 3.3 s in WT, but had a larger contribution (~50%) to the overall Chl

- fluorescence decay. This increase in lifetime and yield of S2QB recombination can be

+ - attributed to a change in the free energy of the primary radical pair P680 PheoD1 to more negative values compared to WT, resulting in an increase of the energetic gap between

199

+ - - - P680 PheoD1 and S2QA (QB ) (Cuni et al., 2004; Rappaport et al., 2005; Cser and Vass,

2007).

The D1-E130L/D2-Q129L double mutant, which combines the effects of the D1-

E130L and D2-Q129L single mutations, had a relatively decreased yield of only 18%

- (72% in WT) for forward ET from QA to QB. However, a faster and more prominent back reaction that contributes to ~58% of the total decay was seen in the D1-E130L/D2-

Q129L double mutant when compared to ~46% in the D1-E130L background strain

(Table 4.3). This suggests that mutagenizing the D2-Q129 residue in the background of

- the D1-E130L mutation also leads to an acceleration and increased yield of S2QB charge recombination asis seen for the D2-Q129 single mutants.

200

Fig. 4.3. Flash-induced Chl fluorescence decay of WT, D2-Q129L, D2-Q129H, D1-

E130L and D1-E130L/D2-Q129L cells measured in the absence of 20 µM DCMU.

201

Strain Fast phase Middle phase Slow phase T1 (ms)/Amp(%) T2 (ms)/Amp(%) T3 (s)/Amp(%)

WT 0.29 ± 0.02/72 ± 3 34.4 ± 19.6/ 17 ± 2 3.30 ± 1.07/11 ± 1 D2-Q129L 0.26 ± 0.02/62 ± 2 34.3 ± 15.5/11 ± 1 0.54 ± 0.07/28 ± 2 D2-Q129H 0.29 ± 0.02/66 ± 2 32.4 ± 15.1/11 ± 1 0.73 ± 0.15/23 ± 2 D1-E130L 0.77 ± 0.91/23 ± 1 98.7 ± 5.3/29 ± 2 98.8 ± 11/48 ± 4 D1-E130L/D2-Q129L 0.51 ± 0.15/18 ± 2 93.5 ± 18.3/24 ± 1 28.1 ± 3.4/58 ± 2

Table 4.3 Chl fluorescence decay kinetics of WT, D2-Q129L, D2-Q129H, D1-E130L and

D1-E130L/D2-Q129L cells in the absence of 20 µM DCMU.

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4.3.4 Chl fluorescence decay in the presence of QB site inhibitors

To assess whether the point mutations at position 129 of the D2 RC protein affected

- - S2QA charge recombination between the S2 state of the OEC and QA , we measured the

Chl fluorescence decay in the presence of a QB site inhibitor. ET inhibitors such as

- DCMU can bind in the QB binding site of PS II and block forward ET from QA to QB.

- Under these conditions, the reoxidation of QA ensues via back reactions with donor side

- components such as the S2 state of the OEC (S2QA charge recombination). Analyzing

Chl fluorescence decay kinetics in the presence of DCMU yields information about changes in the midpoint potentials of cofactors that are energetically placed between the

- OEC and QA in the ET chain. The Chl fluorescence transients measured in the presence of DCMU revealed no significant differences in the D2-Q129L and H mutants relative to

- WT (Fig. 4.4). The decay component representative of S2QA recombination has a lifetime of ~0.26 s in the WT and ~0.23 and 0.27 s in the D2-Q129L and H mutants respectively (Table 4.4). This suggests that the D2-Q129 mutation had no impact on the

- energetic gap between S2 and QA , or on the energetics of any of the intermediate redox cofactors on the active branch.

+ - It is well documented that the free energy of the P680 PheoD1 radical pair is shifted to more negative values in the D1-E130L mutant due to the elimination of the hydrogen bonding interaction to the PheoD1(active) head group (Cuni et al., 2004; Rappaport et al.,

- 2005). Unlike the case of the D2-Q129 mutations where S2QA charge recombination is

- unaffected, the D1-E130L mutation caused a slowing down of S2QA charge recombination as has been observed before (Cuni et al., 2004; Rappaport et al., 2005).

203

- The component representative of S2QA charge recombination in the D1-E130L mutant obtained from the Chl fluorescence decay transient in the presence of DCMU was ~10 s as compared to ~0.26 s in WT. This was expected as mutagenesis of D1-E130L affects the midpoint potential of an ET cofactor (PheoD1) upstream of QB. As seen above, mutagenesis of the D2-Q129 residue to leucine or histidine in the D2-Q129L and D2-

- Q129H single mutants does not alter S2QA recombination. Hence the Chl fluorescence decay of the D1-E130L/D2-Q129L double mutant measured in the presence of DCMU shows no significant differences from the D1-E130L mutant. The mean lifetime of the

- component representative of S2QA charge recombination in the D1-E130L/D2-Q129L mutant was ~11 s, compared to ~10 s in D1-E130L.

From Figs. 4.3 and 4.4 and Tables 4.3 and 4.4, it is clear that mutations of the D2-

Q129 residue impact the kinetics of Chl fluorescence decay only in the absence of QB site inhibitors suggesting the involvement of QB in the changes observed in the Chl fluorescence decay traces of the D2-Q129 mutants compared to WT. More specifically,

- while the rates of forward ET from QA to QB and S2QA charge recombination are

- unchanged in the D2-Q129L and D2-Q129H mutants, the rate of S2QB charge recombination is accelerated 4-6 fold compared to WT. This confirms that mutagenesis of D2-Q129 to leucine or histidine specifically affects the properties of QB while not impacting the properties of any of the active branch cofactors. Moreover, the comparative analysis of Chl fluorescence decay kinetics in the presence and absence of a QB site inhibitor suggests that D2-Q129 mutagenesis induces a decrease in the free energy of the

- S2QB charge pair and in the energetic gap between QA and QB.

204

Fig. 4.4 Chl fluorescence decay of WT, D2-Q129L, D2-Q129H, D1-E130L and D1-

E130L/D2-Q129L cells measured in the presence of 20 µM DCMU. The curves shown were normalized to the same initial amplitude.

205

Strain Fast phase Slow phase

T1 (ms)/Amp(%) T2 (s)/Amp(%)

WT 5.4 ± 2.2/5.4 ± 0.72 0.26 ± 0.01/94.7± 0.72

D2-Q129L 8.7 ± 1.6/7.3 ± 1.3 0.23 ± 0.04/92.7 ± 1.3

D2-Q129H 12.6 ± 3.7/7.0 ± 0.9 0.27 ± 0.02/93.0 ± 0.9

D1-E130L 72.9 ± 32.9/9.8 ± 0.7 10.3 ± 1.6/90.2 ± 0.7

D1-E130L/D2-Q129L 26.4 ± 12.0/15.9 ± 1.4 11.4 ± 2.0/84.1 ± 1.4

Table 4.4 Chl fluorescence decay kinetics of WT, D2-Q129L, D2-Q129H, D1-E130L and

D1-E130L/D2-Q129L cells in the presence of 20 µM DCMU.

206

4.3.5 Chl fluorescence induction kinetics of WT and D2-Q129L, D2-Q129H,

D1-E130L and D1-E130L/D2-Q129L cells

To obtain information on the status of photochemistry in PS II, Chl a fluorescence induction under continuous illumination was measured in WT and mutant strains for up to 1s (Govindjee, 1995). Fig. 4.5 shows the Chl fluorescence induction transients plotted on a logarithmic time scale so that the different rise components could be made more apparent. As reviewed previously, the first phase, labeled I, reflects the net reduction of

- QA to QA (Strasser et al., 1995; Srivastava et al., 1995). The later rise to P is attributed to reduction of the plastoquinone pool. The fluorescence induction kinetics of the WT cells had a normal rise from the basal fluorescence level measured before actinic illumination,

O level, to P. Under the conditions used here, the rise to I (the first peak) occurred in ~30 ms and the total time for reaching the final peak was ~300 ms. The Chl a fluorescence induction transient for the D2-Q129 mutants show similar time constants for reaching the

I and P phases as in WT. However, the relative intensity of the I phase measured at 8 ms after the start of the illumination, increased in the mutants from 0.39 in WT to 0.46 and

0.53 in D2-Q129L and D2-Q129H respectively. This result is consistent with a change in

- - - the equilibrium of the reaction QA QB ↔ QAQB toward QA and/or an increase in the amount of non-QB centers in the mutants (Xiong et al., 1998). Noticeably, and consistent with the effects of the D2-Q129L mutation, the I phase was significantly higher in the

D1-E130L/D2-Q129L mutant with a relative intensity of 0.74 compared to 0.65 in the

- - D1-E130L mutant. Again, this indicated a shift in the equilibrium of QA QB ↔ QAQB

- toward QA .

207

Fig. 4.5 Chl a fluorescence induction transients of WT, D2-Q129L, D2-Q129H, D1-

E130L and D1-E130L/D2-Q129L cells. The data are shown after normalization to the same initial amplitude.

208

4.3.6 Thermoluminescence properties measured in the absence and presence of

20 µM DCMU

Photosynthetic thermoluminescence (TL) refers to photon emission originating from

PS II, stimulated by warming of light-excited samples. It results from recombination of stabilized charge pairs formed during light-driven charge separation. Warming of the sample increases the vibrational energy of the donor and acceptor allowing

* * recombination of the separated charges and re-formation the P680 high energy state. P680 then decays back to the ground state with the emission of a photon (TL) (reviewed in

Inoue, 1996; Vass, 2003; Ducruet and Vass, 2009). The peak position and shape of a TL band is determined by the free energy of activation required for radiative recombination of a particular charge transfer pair. Hence, the properties of TL bands are extremely sensitive to the energetic gaps between donors and acceptors, which in turn are dependent on redox potentials of the recombination partners.

In the absence of ET inhibitors, the TL band generated from WT Chlamydomonas

- cells arises due to charge recombination between the QB and S2 radical pair and has a maximum emission at 28 C (Fig. 4.6). This is commonly referred to as the B band

(Inoue, 1996). The peak temperature of the B band decreased to ~10 and 15 C in the D2-

Q129L and D2-Q129H mutants respectively. This was consistent with the results from the flash-induced Chl fluorescence decay kinetic measurements which revealed a much

- faster rate for S2QB charge recombination in the D2-Q129 mutants than WT. The B band for the D1-E130L mutant was ~10 times more intense than WT, with higher peak temperature of ~36 C (Fig. 4.7). This result was consistent with previous

209 thermoluminescence studies concerning this mutant (Cser and Vass, 2007). However when the D1-E130L mutation was combined with the D2-Q129L mutation in the D1-

E130L/D2-Q129L double mutant, the peak temperature of the B band decreased from

~36 C in D1-E130L to ~18 C in the double mutant (Fig. 4.7). The D1-E130L/D2-

Q129L double mutant manifested the combined effects of the two single mutations

- indicating that the acceleration of S2QB charge recombination, induced by the D2-Q129 mutation, involved the active branch and not the inactive branch cofactors.

Interestingly, the decrease of ~18 C in peak temperature of the B band of the D1-

E130L/D2-Q129L double mutant when compared to the D1-E130L single mutant background is comparable to the decrease of ~18 C in the B band peak temperature of the D2-Q129L single mutant when compared to WT. The decreases in B band peak temperatures induced by mutations of the D2-Q129 residue indicated a decrease in the

- overall stability of the S2QB charge pair. This decrease in stability may be attributed to a change in the redox properties of the S2 state of the OEC or of QB. Considering that the position of the D2-Q129 residue is nowhere close to the OEC in PS II, the more likely explanation is that D2-Q129 mutagenesis affects the redox properties of QB.

210

Fig. 4.6 Thermoluminescence characteristics of the WT and D2-Q129 mutants in the absence of 20 µM DCMU.

211

Fig. 4.7 Thermoluminescence characteristics of the D1-E130L and D1-E130L/D2-Q129L mutants in the absence of 20 µM DCMU.

212

TL data for the WT control and mutants in the presence of ET inhibitors such as

DCMU are shown in Fig. 4.8 and 4.9. DCMU binds in the QB site and impairs forward

- - - ET from QA to QB. This enables the stabilization of QA instead of QB as the negative charge measurable by TL (Rutherford et al., 1982; Inoue, 1996). Under these conditions

- due the lack of QB , the B band is replaced with the so-called Q band which is

- representative of charge recombination between the S2QA charge pair and has a peak emission at ~7 ºC in WT (Fig. 4.8). The Q bands observed in the D2-Q129L and D2-

Q129H mutants had peak temperatures indistinguishable from WT. These results suggest

- that the overall stability of the S2QA charge pair and the stabilization of the S2-state were not affected by mutagenesis of the D2-Q129 residue. This finding also confirms that the modifications of the B band in D2-Q129L, D2-Q129H and the D1-E130L/D2-Q129L double mutant were due to alterations of QB redox potential and not due to changes in the redox properties of the S2-state or any of the other redox components of the active ET branch. Although no changes were apparent in the peak temperature of the Q band in the

D2-Q129 mutants, an increase in Q band intensity was consistently observed. This may

- be attributed to the steady state accumulation of QA in the D2-Q129 mutants relative to

- - WT; indicating a change QA QB ↔ QAQB equilibrium favoring the back reaction in

- - which QA is formed from QB .

As described before, the midpoint potential of PheoD1 and the free energy level of the

+ - primary radical pair P680 PheoD1 is shifted to more negative values in the D1-E130L

+ - mutant than in WT. This causes an increase in the free energy gap between P680 PheoD1

+ - and P680 QA and a slowing down of charge recombination. As shown in Fig. 4.9, the

213 change in primary radical pair energetics was reflected by an increase in peak temperature of the Q band in D1-E130L which occurred at ~25 C when compared with

6-7 C in WT, consistent with previous findings (Rappaport et al., 2005). When the D2-

Q129L mutation was combined with the D1-E130L mutation in the D1-E130L/D2-

Q129L strain, no changes were observed in the peak temperature of the Q band (Fig. 4.9).

This suggests that the D2-Q129L mutation did not induce a change in the free energy of

- the S2QA charge pair in the double mutant when compared to the D1-E130L single mutant background.

The effect of the D2-Q129 mutations on TL properties in the absence of QB site inhibitors (B band) and lack thereof on TL properties in the presence of QB site inhibitors corroborate the Chl fluorescence decay kinetic data. Taken together these data

- conclusively show that mutagenesis of D2-Q129 decreases the stability of S2QB charge

- pair while leaving the energetics of the S2QA state unchanged. Once again, this is indicative that the changes in TL and Chl fluorescence decay seen in the D2-Q129 mutants relative to WT in the absence of ET inhibitors, are due to changes in the redox properties of QB and not due to energetic changes in any of the active branch ET cofactors between the OEC and QA in PS II.

214

Fig. 4.8 Thermoluminescence characteristics of the WT and D2-Q129 mutants in the presence of 20 µM DCMU.

215

Fig. 4.9 Thermoluminescence characteristics of WT, D1-E130L and D1-E130L/D2-

Q129L mutants in the presence of 20 µM DCMU.

216

- 4.3.7 Effect of p-benzoquinone on S2QB charge recombination

To eliminate the possibility that the major TL band observed in the D2-Q129 mutants,

- in the absence of ET inhibitors, arose due to the recombination of S2 with QA (rather than

- QB ), we treated the cells with 100 mM p-benzoquinone (PBQ) that is used to oxidize the

- plastoquinone pool thus minimizing the contribution of charge recombination due to QA

(Rose et al., 2008). Under our conditions, the peak temperature of the B band obtained for WT after PBQ treatment was ~38 C and was attributed to charge recombination

- between the S2/3-states of the OEC and QB . After PBQ treatment, the peak temperature of the B bands in the D2-Q129L and D2-Q129H mutants were 20 C and 25 C, respectively (Fig. 4.10).

After PBQ treatment, the peak temperature of the major TL band (B band) in the D1-

E130L mutant appeared at ~46 C (~38 C in WT). An increase in TL peak temperature

+ - was expected due to an increase in the overall energetic gap between P680 PheoD1 and

+ - P680 QA in the D1-E130L mutant. The introduction of a point mutation at D2-Q129 in the

D1-E130L background as in the D1-E130L/D2-Q129L double mutant decreased the peak temperature of the TL band from ~46 C in the D1-E130L mutant to ~36 C in the double mutant (Fig. 4.11).

217

Fig. 4.10 Thermoluminescence characteristics of WT and D2-Q129 mutants after PBQ treatment.

218

Fig. 4.11 Thermoluminescence characteristics of the D1-E130L and D1-E130L/D2-

Q129L after PBQ treatment.

219

4.3.8 Effects of photoinhibitory light treatment

Based on analysis of the Chl fluorescence decay and TL measurements, it was seen

- that mutations of the D2-Q129 residue decrease the stability of the S2QB charge pair and

- consequently cause an increase in the probability of S2QB charge recombination. It is known that charge recombination reactions in PS II invariably induce photo oxidative

3 1 damage via P680-mediated O2 formation (reviewed in Krieger-Liszkay et al., 2008; Vass and Cser, 2009). Hence to see whether the increased propensity for charge recombination in the D2-Q129 mutants led to an increased susceptibility to photooxidative damage, we subjected thylakoids prepared from the WT and D2-Q129 mutants to photoinhibitory (PI) light treatment and measured the subsequent decrease in oxygen evolving activity over time. It was observed that the D2-Q129L and D2-Q129H mutations caused increased susceptibility to high light (HL) induced damage as was evident from the more rapid decline in oxygen evolving activity of the D2-Q129 mutant thylakoids when compared to

WT (Fig. 4.12).

In order to determine whether the increased susceptibility of the D2-Q129 mutants to photoinhibition had an effect on photoautotrophic growth under high light conditions,

WT and D2-Q129 mutant cells were spotted on HS agar plates at equal cell densities and grown under 500 µmol light m-2 s-1 of white light for two weeks. A slight retardation in growth and increased yellowing of the D2-Q129L and H cultures was observed when compared to WT (Fig. 4.13). This effect on growth in the D2-Q129 mutants was presumably due to an increase in photooxidative damage relative to WT.

220

Fig. 4.12 Residual rates of oxygen evolution measured following photoinhibitory light treatment of WT and D2-Q129 mutant thylakoids.

221

Fig. 4.13 Growth characteristics of the WT and D2-Q129 mutants under high light (500

µmol light m-2 s-1).

222

4.4 Discussion

PS II has parallel C2-symmetry related transmembrane spanning redox cofactor branches, each consisting of two Chls, a Pheo molecule and a plastoquinone. However, only one of these is used during primary charge separation in PS II and is referred to as the active branch. With the exceptions of the plastoquinones, QA and QB, the redox cofactors of the active branch are most closely associated with the amino acid residues of the D1 protein, while the inactive branch cofactors are associated with the D2 protein.

The primary electron acceptor in PS II, PheoD1, shares a crucial hydrogen bonding interaction with the D1-E130 residue such that the abolishment of the hydrogen bond (as in the D1-E130L Chlamydomonas mutant) leads to the near complete impairment of forward ET in PS II. The inactive branch residue analogous to D1-E130 in PS II is D2-

Q129. Analysis of the 2.9 Å PS II crystal structure from cyanobacteria revealed that D2-

Q129 is 3.9 Å away from the PheoD2 head group and hence is unlikely to be a viable hydrogen bond donor to PheoD2. Regardless, in a comparison of all available sequences of the D2 protein, the D2-Q129 residue it is found to be completely conserved unlike the corresponding inactive branch residue in type II BRCs.

In general, little is known about the specific roles of inactive branch amino acid residues in PS II. In this work we explore the effects of replacing D2-Q129, a highly conserved inactive branch acceptor side residue, with a non-conservative amino acid leucine (D2-Q129L) or a conservative amino acid histidine (D2-Q129H) in

Chlamydomonas reinhardtii. For comparison, we also generated the corresponding active branch mutant D1-E130L. Given that the D2-Q129 residue is positioned close to the

223

- inactive branch we wanted to check whether or not S2QB charge recombination in the

D2-Q129 mutants involved any of the inactive branch cofactors. For this, we generated the D1-E130L/D2-Q129L double transgenic strain (or LL mutant) with concurrent mutations in amino acid residues associated with the active and inactive PS II RC cofactor branches. The WT and D2-Q129L, D2-Q129H, D1-E130L and D1-E130L/D2-

Q129L transgenics were examined for their capacity to perform photoautotrophic growth and evolve oxygen. Furthermore, the transgenic strains were studied through Chl fluorescence induction and decay kinetics and thermoluminescence in absence and presence of ET inhibitors. Lastly, the effect of photoinhibitory light treatment on the D2-

Q129 mutants was assessed.

The ability of the D2-Q129L and D2-Q129H strains to carry out photoautotrophic growth on high salt media was comparable to WT under low light (30 µmol light m-2 s-1) conditions, however the D1-E130L and D1-E130L/D2-Q129L mutants grew very poorly compared to WT (Fig. 4.2). The steady-state rates of oxygen evolution measured in mutant cells followed the same trend as seen for the photoautotrophic growth measurement. While the D2-Q129L and D2-Q129H retained up to 70 to 90% of the oxygen evolving activity of WT, the D1-E130L and D1-E130L/D2-Q129L had only ~20 and 10% of WT activity (Table 4.2). This suggested that while the D2-Q129 mutations did not substantially impact forward ET in PS II, the D1-E130L mutation caused a drastic impairment of PS II activity.

The flash-induced Chl fluorescence decay kinetics measured in the absence of

DCMU showed significant differences between the WT and D2-Q129 mutants (Fig. 4.3).

224

Three lifetime components (fast, intermediate and slow) were used to fit the Chl fluorescence decay curves measured in the absence of DCMU. The fast and intermediate components are indicative of the kinetics of forward ET when the QB site is occupied or empty at the time of the flash. While D2-Q129 mutants showed no change in mean lifetimes for forward ET when compared to the WT, the relative contribution of forward

ET to the overall fluorescence decay was decreased compared to WT (Table 4.3). The slow component of the decay curve reflects charge recombination between the S2-state

- - - with QB via the QA QB ↔ QAQB equilibrium. The amplitude of the slow lifetime component increased as a consequence D2-Q129 mutagenesis from 10% in WT to 28% and 23% in the D2-Q129L and D2-Q129H mutants respectively (Table 4.3). In addition,

- the mean lifetime of S2QB charge recombination was accelerated several fold from 3.3 s in WT to 0.54 s and 0.73 s in the D2-Q129L and D2-Q129H mutants respectively. Since

- the rates of forward ET from QA to QB (fast and intermediate lifetimes) are unaffected in

- the D2-Q129 mutants, the acceleration of S2QB charge recombination in the D2-Q129

- - mutants would have to arise from a higher rate of reverse ET between QAQB and QA QB.

Moreover, a larger relative amplitude of the slower kinetic component representative of

- S2QB charge recombination in the D2-Q129 mutants compared to WT could be ascribed to a lower value for the equilibrium constant for sharing an electron between QA and QB or weaker binding of the plastoquinone to the QB site. Since the lifetime of the middle component, indicative of plastoquinone binding in the QB site, did not change in the D2-

Q129 mutants, we can assume that the change in the amplitude is due to a lower equilibrium constant for QA to QB ET. A similar effect is seen in the D1-E130L/D2-

225

- Q129L double mutant in which the rate of the back reaction (S2QB recombination) is accelerated to ~28 s relative to a lifetime of ~100 s in the D1-E130L mutant. Further, the

- relative amplitude associated with S2QB charge recombination also increased from 48% in D1-E130L to 58% in the D1-E130L/D2-Q129L double mutant. Again, we can attribute this increase in amplitude to a lower equilibrium constant for QA to QB ET in the double mutant, induced by the D2-Q129 mutation (Table 4.3).

In the presence of DCMU, when QB is displaced from its binding site and forward ET is blocked, the decay of flash-induced Chl fluorescence proceeds due to the charge

- recombination of QA with S2. No differences were observed in the overall decay kinetics, measured in the presence of DCMU, between WT and the D2-Q129L and H mutants, or between the D1-E130L and D1-E130L/D2-Q129L mutants when compared with each other (Fig. 4.4, Table 4.4). This clearly suggests the involvement of QB in the changes observed in the Chl fluorescence decay kinetics in the absence of DCMU, rather than any of the cofactors upstream of QB in the PS II ET chain.

The redox gap between QA and QB, which determines the driving force for forward

ET, can be estimated from the ratio of the lifetimes of the slow phase in the absence of

- - DCMU (S2QB recombination) and in its presence (S2QA recombination) (Cser et al.,

2008). This calculation yields ~65 mV for the WT which is in very good agreement with previous calculations based on Chl fluorescence decay kinetics (Robinson and Crofts

1983; Allahverdiyeva et al., 2004). By the same calculation, the ΔG(QA-QB) in the D2-

Q129L and D2-Q129H mutants was found to be ~22 mV and ~25 mV respectively. Since

- S2QA recombination and QA redox properties are not affected by D2-Q129 mutagenesis,

226 we can conclude that the decrease in redox gap between QA and QB in the D2-Q129

- mutants occurs due to a lowering of the QB/QB redox potential by ~40-45 mV, bringing it

- closer to the midpoint potential of the QA/QA couple. This decrease in free energy level

- - causes a destabilization of QB and of the S2QB charge separated state, and is responsible

- - - for shifting the QA QB ↔ QAQB equilibrium toward [QA ] in the D2-Q129 mutants.

Thermoluminescence arises from the radiative recombination of charge transfer pairs that recombine in a temperature dependent manner. The peak temperature of a TL band reflects the energy stored in the charge transfer pair, which in turn is dependent on the redox potentials of the individual recombination partners. As explained earlier, TL that arises from the recombination of the S2 state of the OEC on the donor side with the

- - reduced states QA and QB leads to the formation of the Q and B band respectively. The

D2-Q129 mutation induced changes in the peak temperature of only the B band and not the Q band (Table 4.5). This confirmed the results of the Chl fluorescence decay

- measurements that indicated that the overall stability of the S2QB charge pair was

- decreased by the mutations at D2-Q129, while the stability of the S2QA charge pair was unchanged. Hence, results of the TL data supported the suggestion that D2-Q129 mutagenesis specifically impacts the redox properties of QB.

227

Mean Peak Temperature (± 1 °C)

D1-E130L/D2- TL Band WT D2-Q129L D2-Q129H D1-E130L Q129L B Band 28 10 15 36 18

Q Band 7 6 7 25 24

B Band (PBQ 38 20 25 48 38 treatment)

Table 4.5 TL peak temperatures for WT, D2-Q129L, D2-Q129H, D2-E130L and D1-

E130L/D2-Q129L.

228

Substitution of the D2-Q129 residue by biochemically conservative or non-

- - conservative amino acids shifts the semiquinone equilibrium QA QB ↔ QAQB towards

- QA , indicating that the mutagenesis of D2-Q129 has a structural impact localized to the

- - QB site. Evidence for a shift in the QA QB ↔ QAQB equilibrium is obtained from the Chl fluorescence induction kinetics data that show an increased contribution of the I phase in the D2-Q129L, D2-Q129H and D1-E130L/D2-Q129L mutants, which is correlated with

- increased levels of QA after light excitation compared to WT (Fig. 4.5). This has previously been observed in Chlamydomonas mutants D1-R257E and D1-R257M, in

- which a large fraction of QA persists after flash excitation, indicative of an altered

- - - equilibrium constant of the reaction QA QB ↔ QAQB , in the direction of QA (Xiong et

- al., 1998). Further evidence that mutagenesis of D2-Q129 causes a shift in QA QB ↔

- - QAQB (towards QA ) comes from the observation that the intensity of the Q band formed

- due to S2QA recombination increases in the D2-Q129 mutants compared to WT

- suggesting the accumulation of QA after photoexcitation (Fig. 4.8).

Taken as a whole, our experimental data indicate that the mutagenesis of D2-Q129

- causes: 1) a decrease in the free energy gap for recombination of QB with S2, most likely

- due to the lowering in the Em of the QB/QB couple, 2) no significant impact on the redox potential of QA or the donor side components, 3) a shift the equilibrium constant for the

- - - reaction QA QB ↔ QAQB toward QA , and 4) susceptibility to photoinhibitory light due to

- 3 increased S2QB charge recombination and P680-mediated oxidative damage.

Based on the sequence analogy between the D1 and D2 RC proteins, D1-E130, which has a dramatic influence on the midpoint potential of the primary electron acceptor,

229

PheoD1 of the active branch, is comparable to D2-Q129. Therefore, it would be expected the D2-Q129 residue would interact with or hydrogen bond to PheoD2. Based on the results obtained in this study, we cannot rule out the possibility of an effect of D2-Q129 mutagenesis on PheoD2. However, we can conclude that the nature of the amino acid residue in position 129 of the D2 protein is very important in determining the redox potential of QB. While the side chain of D2-Q129 is 3.9 Å away from the PheoD2 head group and is too distant to serve as a hydrogen bond donor; it is located 3.03 Å away from the OH group of the D1-Y254 residue of the D1-de helix. The D1-de helix is situated between the fourth and fifth (D and E) transmembrane regions of the D1 protein and has been shown to interact with QB and its competitive inhibitors (Fig. 4.1 and 4.14)

(Trebst, 1987; Kless et al., 1995). Backbone-dependent in silico mutagenesis of D2-Q129 using the PyMOL 0.99rc6 software (DeLano Scientific LLC, San Carlos, CA) revealed that in silico mutagenesis of this residue causes the predicted distance between the side- chain and the OH group of D1-Y254 to significantly decrease from 3.03 Å in the WT to

1.03 Å and 0.67 Å in the D2-Q129H and D2-Q129L mutants respectively, possibly causing structural interference of the D1-de helix and the QB site (Fig. 4.14).

230

Fig. 4.14. Effect of in silico mutagenesis of the D2-Q129 residue.

231

Mutagenesis of residues located in the de helix of the D1 protein as in the D1-Y254S

- and D1-F255W mutants, often leads to a decrease in stabilization of QB as observed by decreased B band peak temperatures obtained in TL measurements of these mutants

(Ohad and Hirschberg, 1992). Moreover, a study in which combinatorial mutagenesis was applied to a highly conserved portion of the D1-de helix, demonstrated that while many different combinations of amino acids in positions 254 to 257 of the D1 protein satisfied primary PS II function, all the mutants had decreased semiquinone equilibrium

- compared to WT, most likely indicative of a decrease in the QB/QB midpoint redox potential (Kless and Vermaas, 1995). More evidence for the involvement of the D1-de

- helix in determining QB redox potential and hence the stabilization of the S2QB charge pair comes from the site-directed mutagenesis of the highly conserved residue located

D1-R257 residue located at the C-terminal end of the de helix. While this residue points away from the QB binding pocket into the stromal surface, site-directed mutants such as the D1-R257E,K,Q transgenics have Chl fluorescence decay and TL properties

- - suggestive of a lowered QB redox potential and a shifted QA QB ↔ QAQB equilibrium

(Xiong et al., 1998; Rose et al., 2008).

Hence combining the Chl fluorescence, TL and in silico mutagenesis results obtained in our study, we can hypothesize that the mutagenesis of an ―inactive branch‖ residue

D2-Q129 (that albeit resides close to the head group of PheoD2) has an indirect consequence on the redox potential of QB caused via interactions with the D1-de helix, which is known to play a role in determining the redox potential of QB. It is noteworthy that D2-F252 is the QA site residue of the D2-de helix that corresponds to D1-Y254 of the

232

D1-de helix (Fig. 4.1). Unlike D1-Y254, the side chain of D2-F252 lacks the additional

OH group which possibly minimizes steric interference between amino acid residues close to the PheoD1 head group and the side chains of the D2-de helix residues in the QA site.

This study provides insight into the extent of asymmetry between the two ET branches in PS II where analogous active and inactive branch mutations can impact non- analogous cofactors of the two branches. Our experimental data elucidate further the impact of individual inactive branch residues on PS II function. Moreover, this is one of few studies, which have demonstrated the role of an inactive branch residue in determining the redox properties of a cofactor that takes part in primary ET in PS II. Our results also provide the first report of a single amino acid substitution of the D2 protein that affects the redox properties of QB. Lastly, this study includes the first instance of simultaneous mutagenesis of active and inactive branch residues and of the D1 and D2 proteins, in PS II.

233

Chapter 5

Summary

The recent impetus toward finding alternative sources of renewable and clean fuels to meet our growing energy demands and to mitigate the environmental impacts of fossil fuel combustion has directed renewed interest in the potential benefits of algae as a biofuel feedstock. Microalgae have several advantages over other biofuel crops because they: i) have faster growth rates ii) higher photon conversion efficiencies iii) can be harvested year-round and iv) have the potential for reduced dependence on fresh water.

The productivity (biomass yield) of algal cultures and biofuel production is closely correlated with the efficiency of photosynthesis (Fig. 5.1) (Schenk et al., 2008; Dismukes et al., 2008).

During photosynthesis, eukaryotic photosynthetic organisms like green algae and plants use light energy to drive the oxidation of water and the synthesis of ATP and

NADPH that are eventually used in the production of sugar. Light energy is harvested by the peripheral light harvesting complexes or antennae, LHCI and LHCII of the two photosystems, PS I and PS II, and transferred to the central reaction center (RC) complexes where primary photochemistry occurs. The efficiency of photosynthesis is

234 dependent both on the light-harvesting capacity of PS II well as on the properties of the

PS II RC redox active cofactors that carry out photosynthetic electron transfer.

Given the importance of photosynthesis and photosystem II to the production of biofuels we carried out a detailed study of the photosystem II complex in which we investigated the impacts of altering the light-harvesting capacities of the peripheral PS II antennae and the efficiencies of forward and back electron transfer in PS II RCs, on the overall efficiency of photosynthesis and photoautotrophic growth productivity. The specific aims of the work presented in this thesis were: 1) to develop genetic strategies to modulate PS II peripheral antennae size with the goal of maximizing photosynthetic productivity (and biomass yield) under all light environments and 2) to study the effects of site-specific mutagenesis of the PS II RC polypeptides (D1 and D2) that impact the properties of the primary donor (ChlD1) and terminal acceptor (QB) of PS II, on the yield of forward electron transfer (ET) and charge recombination and on photosynthetic productivity. We employed the unicellular green alga Chlamydomonas reinhardtii as a model system for our studies on ―the modulation of energy and electron transfer processes of PS II‖.

235

Fig. 5.1 Photosynthesis and biofuel production (Schenk et al., 2008, Bioenerg. Res. 1: 20

– 43).

236

Chapter 2 provides an overview of the effects of altering PS II peripheral antenna size on photosynthetic productivity and growth of C. reinhardtii. Low light acclimated

Chlamydomonas cells typically possess larger light harvesting antennae so as to maximize light capture at limiting light conditions. However a negative consequence of having very efficient light harvesting complexes is that photosynthetic electron transfer in nearly all photosynthetic eukaryotes reaches becomes light saturated at only 25% of full sunlight intensity (Polle et al., 2002). Under high light conditions, the over-excitation of the light-harvesting antenna leads to the induction of short-term photoprotection responses (NPQ) that can dissipate up to ~80% of the absorbed photons as heat or fluorescence to minimize 3Chl* induced photooxidative damage, causing large decreases in light utilization and photosynthetic productivities (Polle et al., 2002). Algae that are defective in Chl b synthesis due to the lack of the enzyme, chlorophyllide a oxygenase

(CAO), have smaller PS II light harvesting antennae and do not light saturate electron transfer. However, they are less efficient at harvesting photons at lower light intensities.

To increase the absorption and utilization of light at both low and high light intensities, we used an RNAi approach to down-regulate the expression of the CAO gene and generate transgenic algae (CR transgenics) with intermediate levels of Chl b and PS

II peripheral antennae sizes. The CR transgenics had 1.5-2.2 fold higher ratios of Chl a/b, which are an indicator of preferentially decreased PS II antenna size. We demonstrated that modulating the levels of Chl b, which binds preferentially to the PS II peripheral antenna, was sufficient to affect the size of the PS II-LHCII antenna complex and also to substantially alter biomass yields. Chl fluorescence induction kinetics of the CR

237 transgenics showed that they light saturated ~10-15% more slowly than the wild-type.

We also confirmed a linear but inverse relationship between the Chl a/b ratio and the size of the PS II peripheral antenna. Decreased PS II antenna size translated to a ~2 fold increase in light saturated rates of photosynthesis for the CR transgenics compared to wild-type on a per Chl basis and a ~1.6 fold increase on a per cell basis. Additionally, the

CR transgenics with intermediate PS II antennae sizes outperformed (30% higher growth) wild-type cells in monocultures at light saturating intensities without an impairment in effective photosynthesis or growth rates at sub-saturating light intensities.

Over the course of a year, light intensities and durations are expected to vary dramatically. It has been hypothesized that dynamic modulation of PS II antennae size can optimize light harvesting efficiency at both low and high light intensities. In this study, we demonstrated that the modulation of PS II antennae size by light-regulated control of Chl b accumulation may provide an efficient strategy to optimize PS II antennae size to seasonal changes in light intensity. Using a 13 bp NAB1 binding site fused to the 5‘ end of the CAO transcript, we induced the post-transcriptional regulation of CAO and Chl b accumulation and hence of PS II antenna size. NAB1 is a RNA- binding protein whose binding capacity is light regulated and redox sensitive. Under high light intensities NAB1 binds to its RNA-binding site (NAB1 binding site, N1BS) on

LHCII mRNAs, sequestering them into translationally silent messenger ribonucleoprotein thereby negatively regulating their translation and causing a reduction of LHCII content under high light (Mussgnug et al., 2005, Wobbe et al., 2009). In this study, we fused the

NAB1 binding site to the CAO gene to control its expression and therefore Chl b

238 accumulation in a light dependent manner. At high light intensities the NAB1 protein binds to its respective mRNA binding site on the engineered CAO transcript, repressing its translation and the synthesis of Chl b, resulting in a reduced PS II peripheral antenna size. The reverse scenario is expected at low light intensities, when NAB1 does not bind to its target mRNA and allows its translation to occur (Fig. 5.2).

Our results from three independent transformants (N1BS-CAO-4, 7, and 77) expressing the modified CAO gene showed that Chl a/b ratios increased under high light acclimation resulting in reduced PS II antenna size. Conversely, a decrease in Chl a/b ratios were observed under conditions of low irradiances which were indicative of an increase in PS II antenna size. To support these interpretations we carried out flash induced Chl fluorescence induced kinetics of low light and high light grown cultures. The

N1BS-CAO-4,7 and 77 strains exhibited up to ~10% increase in the induction of light saturated Chl fluorescence compared to the complemented wild-type CAO-4 under low light intensities, and a ~10% reduction in Chl fluorescence yield relative to 90% light saturation yield for wild type when grown under high light conditions. This light- dependent change in antennae size in the transgenics was substantially greater than that observed in wild-type cells and was seen to occur on the order of days. In our experiments, Chlamydomonas cells were light acclimated for six day periods during which the NAB1-regulated (light-dependent) changes in Chl b accumulation and PS II antennae size occurred. Hence this strategy would be applicable in maximizing photon absorption and utilization during seasonal changes in light intensity.

239

Fig. 5.2 Results summary of the NAB1-CAO transgenics characterized in Chapter 2.

240

Chapter 3 provides an in depth analysis of the effects of site-directed mutagenesis of the protein environment of the primary electron donor in PS II, ChlD1, and its impact on photosynthetic productivity and electron transfer. In the type II bacterial RCs, the primary electron donor is the BChl special pair (BChlSP), a dimer of BChl molecules (reviewed in

Kirmaier and Holten, 1987). However it has been shown that in PS II, the active branch monomeric Chl, ChlD1, and not the Chl special pair (ChlSP), is the more plausible primary electron donor and the site of primary charge separation in PS II (Groot et al., 2005;

Holzwarth et al., 2006). We modified the local protein environment of ChlD1 to elucidate its function charge separation as well as to study the effects of the proximal amino acid residues on its redox properties. The closest residue to ChlD1 is D1-T179 and is found overlying the coordinated Mg2+ but is not close enough to directly serve as a ligand to it

(Ferreira et al., 2004; Loll et al., 2005). Axial coordination to ChlD1 is thought to be provided by a water molecule hydrogen bonded to D1-T179O. Despite not being involved directly in ligating ChlD1, D1-T179 is completely conserved throughout oxygenic photosynthesis.

We replaced the D1-T179 residue with serine, aspartate, asparagine, histidine (more polar residues) and isoleucine to generate the D1-T179S, D, N, H and I Chlamydomonas

PS II mutants respectively. We observed that mutagenesis of this residue impairs photoautotrophic growth and photosynthetic oxygen evolution where the D1-T179S, D,

H, N and I mutants had 71%, 64%, 60%, 43% and 37% of the oxygen evolution activity of the wild-type control. The Chl fluorescence value of F0 which reflects the Chl fluorescence emanating from the PS II antenna in competition with excitation energy

241 transfer to the RC, were elevated in the D1-T179N and I mutants, indicative of inefficient excitation energy transfer from the proximal PS II antenna complex to the RC core.

Interestingly, we observed a correlation between the predicted increase in proximity of the mutagenized amino acid residue to ChlD1 and F0, suggesting that the efficiency of excitation energy transfer to the RC Chls from the proximal PS II antenna was compromised with decreasing predicted coordination distance.

Mutagenesis of the D1-T179 residue caused altered flash-induced Chl fluorescence decay kinetics relative to wild-type. The slow kinetic component of the Chl fluorescence decay obtained in the absence of a QB site ET inhibitor is representative of charge

- recombination between the S2 state of the OEC and QB . In the wild-type, the lifetime of

- S2QB charge recombination is ~3.1 s. The lifetime of this component is accelerated to

- 1.3, 1.0 and 0.9 s in the D1-T179D, N and H mutants respectively. S2QB charge recombination in the D1-T179S transgenic was most similar to wild-type with a lifetime

- of ~2.2 s. Finally, the S2QB back reaction in the D1-T179I mutant was slower, occurring in ~5.6 s. The same trend was observed in Chl fluorescence decay kinetics in the presence of QB site ET inhibitors in which the slow kinetic component is attributed to

- - S2QA charge recombination. The lifetime component for S2QA charge recombination was 0.25 s in the wild type (D1-T179) which was accelerated to 0.22, 0.12, 0.09 and 0.07 s in the D1-T179S, N, D and H mutants respectively (Fig. 5.3). In the D1-T179I mutant

- however the rate of S2QA charge recombination is slowed to 0.36 s. The much faster

- - rates of S2QA and S2QB charge recombination in the D1-T19N, D and H mutants were

+ - explained by a decrease in free energy difference of the primary radical pair P680 PheoD1

242

- - and the S2QA or S2QB states respectively, compared to wild-type. Conversely, the slower rates of charge recombination in the D1-T179I mutant was attributed to an increase in the free energy gap between the primary radical pair and the secondary charge separated states.

- Thermoluminescence (TL) measured in the absence of DCMU arises due to S2QB recombination and yields a characteristic band (B band) with a peak emission at ~28 C in the wild type. TL arises from the radiative recombination of stabilized charge pairs

1 * formed during light-driven charge separation giving rise to the P680 Phe state, which decays via the emission of a photon (TL). The peak position and shape of a TL band is determined by the free energy of activation required for radiative recombination of a particular charge transfer pair. The peak temperature and intensity of the B band decreased to ~21, 20, and 19 C in the D1-T179D, H and N mutants respectively relative

- to 28 C in wild-type. The changes in the free energy of S2QB recombination in the D1-

T179D, H and N mutants relative to the wild-type were calculated using the normalized

- intensity of the TL bands and yielded an overall decrease in the free energy for S2QB

* + - recombination and an increase in ΔG(P680 ↔P680 PheoD1 ) of ~16 to 25 meV compared to wild-type. In the presence of DCMU, QB is displaced from the QB site and the dominant

- TL band arises due to S2QA charge recombination. In the wild-type the peak temperature of this band was ~5-6 C mutants. The peak temperature of the Q band also decreased in the D1-T179D, H and N mutants appearing between 0-3 C with a corresponding decrease in intensity. The D1-T179S mutant had similar TL characteristics as the wild-

243 type while the Q band in the D1-T179I mutant had a slightly higher peak temperature of

8 C but less than ~10% of wild-type TL intensity.

The combined results of the thermoluminescence and flash-induced Chl fluorescence decay measurements suggested that mutagenesis of D1-T179 to amino acids more polar

+ than threonine, cause a shift in the Em of P680/P680 to more positive values such that the probability of indirect radiative charge recombination decreases due to an increase in the

* + - energetic gap between P680 and the primary radical pair P680 PheoD1 . However, the

- overall rate for S2QA charge recombination was accelerated due to a decrease in the

- energetic gap between the primary radical pair and the S2QA state. The effects of the mutagenesis of D1-T179, on the redox properties of P680 provide support to the suggestion that ChlD1 is the primary electron donor in PS II. We also concluded that the properties of the D1-T179 residue are important in determining the efficiencies of excitation energy transfer and of forward ET and charge recombination in PS II.

Therefore, in contrast to type II bacterial reaction centers (BRC) (in which a histidine resides in the position analogous to D1-T179 in PS II) having a histidine residue in closest proximity to the ChlD1 monomer of the active branch reduces the efficiency of forward ET and oxygenic photosynthesis. From an evolutionary perspective while the PS

II RC maintains many features of its BRC progenitor, it is foreseeable that the residue near the monomeric ChlD1 was substituted (with threonine) during the evolution of oxygenic photosynthesis, to provide for an efficient balance between the ability to generate a highly oxidizing primary donor cation for water oxidation, and forward and back electron transfer.

244

Fig. 5.3 Summary of important results of D1-T179 mutagenesis described in Chapter 3.

245

In Chapter 4, we explored the extent of asymmetry between the two pseudo- symmetrical cofactor branches of PS II of which only one, the active branch, takes part in primary ET. The primary electron acceptor in PS II is the active branch pheophytin,

PheoD1 (Klimov et al., 1977; Klimov and Kravnoski, 1981; Klimov, 2003; Groot et al.,

2005; Holzwarth et al., 2005). Similar to the purple BRCs there are two symmetry-related pheophytins (PheoD1 and PheoD2) in PS II. The redox properties of PheoD1 are modulated by the nature of the amino acid residue in position 130 of the D1 protein. In wild-type chloroplastic PS II, the D1-130 residue is a glutamate and it serves as a hydrogen bond donor to the PheoD1 head group (Dorlet et al., 2001). It has been shown that mutagenesis of D1-130 to amino acids that weakened the hydrogen bonding interaction to PheoD1

- shifted the midpoint potential of the PheoD1/PheoD1 couple to more negative values, making forward ET transfer from PheoD1 to QA less probable (Cuni et al., 2004;

Rappaportet al.,2005; Cser and Vass, 2007). Hence, the D1-E130L PS II mutation effectively resulted in an impairment of forward ET and a consequent reduction in primary charge separation yield. The inactive branch residue analogous to D1-E130 in PS

II is D2-Q129. D2-Q129 was also previously suggested as being a possible hydrogen bond donor to the head group of PheoD2 (Xiong et al., 1996; Kern and Renger, 2007).

However, an analysis of the 2.9 Å PS II crystal structure from the cyanobacterium

Thermosynechococcus elongatus showed that the side chain of D2-Q129 is 3.9 Å away from the PheoD2 head group and therefore may not be in close enough proximity to serve as a viable hydrogen bond donor. A comparison of all 152 protein sequences available for

246 the D2 polypeptide of PS II from a variety of oxygenic photoautotrophs showed that D2-

Q129 is completely conserved across all sequences.

In general, we know little about the specific roles of acceptor-side inactive branch residues in PS II. In this study, we elucidated the role of the highly conserved acceptor side residue on the inactive branch, D2-Q129. In contrast to the well understood role of its active branch counterpart (D1-E130), no previous function had been assigned to D2-

Q129 based on experimental data. We employed a mutagenesis approach in addressing the possible roles of this residue. D2-Q129 was replaced by a non-conservative hydrophobic leucine residue (D2-Q129L) or a conservative histidine residue (D2-Q129H) in Chlamydomonas reinhardtii. We also generated the analogous D1-E130L single mutant as well as the D1-E130L/D2-Q129L double mutant for simultaneous comparison of the effects of the analogous active and inactive branch residues. The resulting transgenics were analyzed by their photoautotrophic growth characteristics, oxygen evolution rates, Chl fluorescence induction and decay kinetics, thermoluminescence properties and sensitivity to photoinhibitory light treatment.

The photoautotrophic growth of the D2-Q129L and D2-Q129H single mutants was comparable to the complemented wild-type under low light intensities and had ~70% and

90% of wild-type oxygen evolving activity respectively. The D1-E130L mutant that is known to be impaired in photosynthesis and the D1-E130L/D2-Q129L double mutant had poor photoautotrophic growth and only ~20% and 10% of wild-type oxygen evolution activity, respectively.

247

The Chl fluorescence decay kinetics of the D2-Q129L and D2-Q129H mutants in the absence of the QB site inhibitor DCMU revealed a decrease in the total contribution of forward ET to the overall Chl fluorescence decay from 90% of the total in the wild-type to ~73 and 77% of the total in the D2-Q129L and D2-Q129H mutants respectively.

- However, the mean lifetime for forward ET from QA to QB was not affected by the D2-

Q129 mutation. The contribution of the slow component, representative of charge

- recombination from QB to the S2 state was increased from ~10% in the wild-type to ~28 and 23% in the D2-Q129L and H mutants, respectively. Furthermore, the lifetime

- component for S2QB charge recombination was accelerated ~4 to 6 fold from ~3.3 s in wild-type, to 0.54 and 0.73 s in the D2-Q129L and H mutants, respectively. The Chl fluorescence decay kinetics obtained for the D1-E130L mutant was similar to previously

- published results in which a large increase in the lifetime of S2QB recombination was observed. This has been attributed to a change in the free energy of the primary radical

+ - pair P680 PheoD1 to more negative values compared to wild-type, resulting in an increase

+ - - - of the energetic gap between P680 PheoD1 and S2QA (S2QB ) and a slowing down of electron transfer (Cuni et al., 2004; Rappaport et al., 2005; Cser and Vass, 2007). The mutagenesis of the D2-Q129 residue in the background of the D1-E130L mutation as in the D1-E130L/D2-Q129L double mutant, led to a faster back reaction with a larger associated amplitude which contributed to ~58% of the total decay when compared to

~46% in the D1-E130L background strain. This suggested that mutagenizing the D2-

Q129 residue in the background of the D1-E130L mutation led to an acceleration and

- increase in yield of S2QB charge recombination as was seen for the D2-Q129 single

248 mutants. Interestingly no differences were observed in the Chl fluorescence decay kinetics between the wild-type and D2-Q129 mutants (or between the D1-E130L and D1-

E130L/D2-Q129L mutants) in the presence of a QB site inhibitor DCMU indicating that

- the lifetime for S2QA charge recombination was not altered due to D2-Q129 mutagenesis.

This result confirmed that mutagenesis of D2-Q129 to leucine or histidine specifically affected the properties of QB while not impacting the properties of any of the active branch cofactors. Moreover, the comparative analysis of Chl fluorescence decay kinetics in the presence and absence of DCMU suggested that D2-Q129 mutagenesis resulted in a

- decrease in the free energy of the S2QB charge pair and in the energetic gap between QA and QB.

The redox gap between QA and QB, which determines the driving force for forward

- - ET, was estimated from the ratio of the lifetimes of S2QB and S2QA recombination. This calculation yielded a change in the free energy difference of ~65 mV for ΔG(QA-QB) in wild-type whereas ~22 mV and ~25 mV were obtained for the D2-Q129L and D2-Q129H

- mutants, respectively. Since S2QA recombination and QA redox properties were not affected by D2-Q129 mutagenesis, we concluded that the decrease in redox gap between

- QA and QB in the D2-Q129 mutants occurred due to a lowering of the QB/QB redox potential by ~40-45 mV. This decrease in the free energy gap between QA and QB in the

- - D2-Q129 mutants, caused a destabilization of QB and of the S2QB charge separated state,

- - which was inferred as being responsible for shifting the QA QB ↔ QAQB equilibrium

- toward [QA ], relative to wild-type.

249

The peak temperature of a TL band reflects the energy stored in the charge transfer pair, which in turn is dependent on the redox potentials of the individual recombination partners. As explained earlier, TL that arises from the recombination of the S2 state of the

- - OEC on the donor side with the reduced states QA and QB leads to the formation of the

Q and B band respectively. Consistent with the Chl fluorescence decay kinetics in which differences between the D2-Q129 mutants and wild-type were observed only in the absence of DCMU, the D2-Q129 mutation induced changes in the peak temperature of only the B band (measured in the absence of DCMU) and not the Q band (measured in the presence of DCMU) (Fig. 5.4). This result confirmed that the overall stability of the

- S2QB charge pair was decreased by the mutations at D2-Q129, while the stability of the

- S2QA charge pair was unchanged. Hence, the results of our TL data supported the hypothesis that D2-Q129 mutagenesis specifically impacts the redox properties of QB.

Based on the results described here, we conclude that the nature of the amino acid residue in position 129 of the D2 protein is very important in determining the redox potential of QB. Interestingly, D2-Q129 is located 3.03 Å away from the OH group of the

D1-Y254 residue of the D1-de helix. Backbone-dependent in silico mutagenesis of the

D2-Q129 residue revealed that the predicted distance between the side-chain and the OH group of D1-Y254 and the D2-Q129 side chain significantly decreased from 3.03 Å in the wild-type to 1.03 Å and 0.67 Å in the D2-Q129H and D2-Q129L mutants, respectively. The predicted decrease in distance as a result of D2-Q129 mutagenesis was thought to cause structural changes in the D1-de helix and consequently in the QB site.

250

Mutations of D1-de helix have previously been shown to cause decreased stabilization of

- QB (Ohad and Hirschberg, 1992).

- The increased yield of S2QB charge recombination in the D2-Q129 mutants relative to wild-type also led to an increase in susceptibility to photoinhibitory light presumably

3 due to P680-mediated oxidative damage (reviewed in Krieger-Liszkay et al., 2008). It was observed that the oxygen evolution activity of thylakoids isolated from the D2-Q129L and D2-Q129H mutants decreased more rapidly under photoinhibitory light treatment relative to the wild-type. Further, a slight retardation in growth and increased yellowing of the D2-Q129L and D2-Q129H cultures was observed under high light intensities (500

µmol light m-2 s-1) when compared to wild-type implying increased levels of high light- induced photooxidative damage in the D2-Q129 mutants.

Hence, based on our experimental data we concluded that the mutagenesis of D2-

- Q129 causes a decrease in the free energy gap for recombination of QB with S2, most

- likely due to the lowering in the Em of the QB/QB couple while not inducing any noticeable impact on the redox potential of QA or the donor side components. Further,

- - D2-Q129 mutagenesis induced a shift in the equilibrium of the reaction QA QB ↔ QAQB

- - toward QA and increased susceptibility to photoinhibitory light due to increased S2QB

3 charge recombination and P680-mediated oxidative damage. The experimental data obtained here increase our understanding of the specific impact of individual inactive branch amino acid residues on electron transfer in PS II and of the structural asymmetry evident in PS II. Moreover, this is one of few studies that has clearly demonstrated the

251 role of an inactive branch residue in determining the redox properties of a cofactor that takes part in primary ET in PS II.

252

Fig. 5.4 Results summary of Chapter 4.

253

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