Regulation of Photosynthesis in plants under abiotic stress
A thesis submitted to the University of Manchester for the degree of PhD in the Faculty of Life Sciences
2014
Sashila Abeykoon Walawwe
1 Table of contents
List of Figures 5 List of Tables 9 Abbreviations 10 Abstract 12 Declaration 13 Copyright Statement 13 Acknowledgements 14
Chapter 1- General Introduction 15 1.1. Introduction 16 1.2. Photosynthesis 18 1.2.1. Light capture and electron transport chain 18 1.2.2. Cyclic electron transport 31 1.2.3. The Calvin-Benson-Bassham cycle 39 1.3. Effects of abiotic or environmental stress on plants 43 1.3.1. Salt stress 43 1.3.1.1. Effects of salt stress on plants 45 1) Effects of salt on plant growth 45 2) Effects of salt on photosynthesis 46 3) Effects of salt on water relations and ion balance in plants 47 4) Effects of salt on photosynthetic pigments, proteins and lipid 48 composition 5) Effects of salt on leaf anatomy and the structure of chloroplast 49 6) Effects of salt on the nitrate and malate metabolism 50 1.3.1.2. Sensing and signal transduction in salt stress tolerance 51 1.3.1.3. Transcriptomics and proteomics of salt tolerance 55 1.3.1.4. Salt tolerance mechanisms in plants 59 1) Ion regulation and compartmentalization 59 2) Accumulation of compatible solutes 64 3) Involvement of the antioxidant enzymes 65 4) Involvement of plant hormones 67 1.3.1.5. Effects of salt stress on other photosynthetic organisms 69
2 1) Cyanobacteria 69 2) Algae 73 1.3.2. Drought stress 75 1.3.3. Heat stress 76 1.3.4. Low temperature stress 77 1.4. Effects of Environmental stress on the electron transport of photosynthesis 80 1.4.1. Effects on energy use in photosynthesis 81 1.4.1.1. Reactive oxygen species (ROS) formation 82 1.4.2. Effects on components of the electron transport 85 1.5. Regulation of electron transport chain of photosynthesis under stress conditions 89 1.5.1. State-transitions (qT) 89 1.5.2. High-energy state Quenching (qE) 92 1.5.3. Other electron transport pathways involved in regulatory process 98 1.5.3.1. Mehler Reaction 98 1.5.3.2. Chlororespiration 100 1.6. Involvement of Plastid terminal oxidase (PTOX) in alternative electron transport 103 in the electron transport chain of Thellungiella salsuginea 1.7. Aims and Objectives 110
Chapter 2- Effect of salt stress on the regulation of photosynthesis in barley 112 (Hordeum vulgare L.)
Preface 113
2.1. Abstract 114 2.2. Introduction 115 2.3. Materials and Methods 119 2.4. Results 130 2.5. Discussion 140
Chapter 3- Physiological evaluation of salinity stress in two rice varieties from Sri Lanka 149 Preface 150 3.1. Abstract 151 3.2. Introduction 152 3.3. Materials and Methods 157
3 3.4. Results 169 3.5. Discussion 187
Chapter 4- Regulation of photosynthesis in Thellungiella salsuginea under abiotic stress 197
Preface 198 4.1. Abstract 199 4.2. Introduction 200 4.3. Materials and Methods 206 4.4. Results 215 4.5. Discussion 231
Chapter 5- General Discussion 239
Bibliography 247
Word Count: 59755
4 List of Figures
Chapter 1- Introduction
Figure 1.1. Schematic model of the major protein complexes involved in electron transport 20 chain in photosynthesis Figure 1.2. Schematic model representing the proteins of light harvesting complex II 23 (LHCII) and reaction centre core (RCII) of photosystem II (PSII) Figure 1.3. The model of Q cycle representing the electron and proton transport in the 26
cytochrome b6f Figure 1.4. Schematic model representing the proteins of light harvesting complex I (LHCI) 28 and reaction centre core (RCI) of photosystem I (PSI) Figure 1.5. Schematic model representing the chloroplast ATP synthase 30 Figure 1.6. Possible pathways of cyclic electron transport around PSI 38 Figure 1.7. A diagram representing the major steps in the Calvin cycle or the light-independent 42 reactions Figure 1.8. Cellular Na+ transport mechanisms and important components of the salt 54 stress responses in plant root cells Figure 1.9. Regulation of electron transport through qE 97
Chapter 2
Figure 2.1. Far-red light induced signal giving the 100% of P700 122
Figure 2.2. Typical fluorescence signal showing all the reference points 123
Figure 2.3. P700 oxidation of which is induced by the actinic light was measured during 126 a 100 milliseconds period of darkness Figure 2.4. The relative concentration of 'active' PSI centres (centres that can be oxidized by 127 light and are then rapidly re-reduced during a period of darkness)
2 Figure 2.5. The first leaf of barley plant showing the section of approximately 2.1 cm 128 (length 3 cm x width 0.7 cm between 4 cm and 7 cm from the leaf tip of each leaf) area Figure 2.6. Gas exchange parameters of barley plants subjected to varying degrees 132 of salinity
5 Figure 2.7. Maximum quantum yield (Fv/Fm) of control and salt-treated barley plants 133
Figure 2.8. The effect of salt treatment on the efficiency of PSII (ΦPSII), relative 135 electron transport of PSII (PSII ETR) and non-photochemical quenching (NPQ) of barley plants
Figure 2.9. The effect of salt treatment on redox state of P700, relative proportion of the 138 active PSI centres, rate constant for P700 reduction and electron transport rate of PSI (PSI ETR) of barley plants
Figure 2.10. The effect of salt treatment on the leaf chlorophyll content per leaf area and 139 chlorophyll a/b ratio in barley
Chapter 3
Figure 3.1. Far-red light induced signal giving 100% of P700 161
Figure 3.2. Typical fluorescence signal showing all the reference points 162
Figure 3.3. P700 oxidation induced by the actinic light was measured during a 164 100 milliseconds period of darkness
Figure 3.4. The relative concentration of 'active' PSI centres (centres that can be oxidized by 165 light and are then rapidly re-reduced during a period of darkness)
Figure 3.5. Fluorescence signal showing the relaxation kinetics 167
Figure 3.6. 34 days old At-354 and Bg-352 control and salt treated plants at the early 170 vegetative stage
Figure 3.7. Change in leaf area of two rice varieties At-354 and Bg-352 at the early 171 vegetative stage and at the flowering stage when exposed to 50 and 100 mM of NaCl
Figure 3.8. The effect of salinity on leaf chlorophyll content per leaf area and chlorophyll 173 a/b ratio in two varieties of rice, At-354 and Bg-352
Figure 3.9. Gas exchange parameters measured in rice plants at the early vegetative stage 175 and the flowering stage of Bg-352 and At-354 exposed to: 0, 50 and 100 mM
6 of NaCl
Figure 3.10. CO2 assimilation rate (A) as a function of internal CO2 concentration (Ci) in 176 plants at the tillering stage and flowering stage of Bg-352 and At-354 exposed to: 0, 50 and 100 mM of NaCl
Figure 3.11. Maximum quantum yield (Fv/Fm) of salt stressed (50 or 100 mM) and control 177 plants at early vegetative stage and at flowering stage of At-354 and Bg-352
Figure 3.12. Photochemical efficiency (ΦPSII) and relative linear electron transport rate of 179 PSII (PSII ETR) plants at the early vegetative stage and the flowering stage of Bg-352 and At-354 exposed to: 0, 50 and 100 mM of NaCl
Figure 3.13. Non Photochemical Quenching (NPQ) plants at the early vegetative stage and 181 the flowering stage of Bg-352 and At-354 exposed to: 0, 50 and 100 mM of NaCl
Figure 3.14. Chlorophyll fluorescence relaxation kinetics of control and salt treated plants at 182 the flowering stage recorded using the PAM 101 fluorometer.
Figure 3.15. Redox state of P700 and the proportion of 'active' PSI centres of two rice 184 varieties subjected to different salt concentrations plants at the early vegetative stage and the flowering stage
Figure 3.16. The rate constant of P700 reduction and PSI electron transport rate (PSI ETR) of 186 plants at the early vegetative stage and the flowering stage of Bg-352 and At-354 exposed to: 0, 50 and 100 mM of NaCl.
Chapter 4
Figure 4.1. Images showing the physical changes of leaves of 7-week old T. salsuginea 216 when exposed to stresses.
Figure 4.2. Immunoblots and relative band intensity of PTOX, cytochrome f (Cyt f) and 218 PsbA from control, salt-treated, droughted and plants exposed to different growth
7 irradiance
Figure 4.3. PTOX gene expression along with actin and the relative expression level of 220 PTOX mRNA in control and salt-treated plants.
Figure 4.4. Change in the efficiency of PSII (ΦPSII) measured in control and stressed 222
-1 plants at the different light intensities and CO2 concentration of >1200 μL L
Figure 4.5. Change in the electron transport rate of PSII (ETR of PSII) measured in control 223
plants and stressed plants at the different light intensities and CO2 concentration of >1200 μLL-1
Figure 4.6. Blue-native PAGE showing the separated complexes of isolated thylakoids of T. 225 salsuginea
Figure 4.7. PTOX genomic sequence of T. salsuginea including exons, introns and 229 untranslated regions (UTRs) coding sequence of PTOX which indicates the annealing sites of primers used in rt-PCR analysis.
Figure 4.8. The schematic representation of the genomic structure of PTOX contains exons, 230 introns and transcription start site (ATG) and stop codon (TAA)
Chapter 5- General Discussion
8 List of Tables
Chapter 1
Table 1.1. Functional groups of genes/proteins induce under salt stress 56
Table 1.2. Selective examples of genes/proteins induced by salt stress 57
Table 1.3. Reactions of the Mehler or water-water cycle 99
Chapter 3
Table 3.1. Fast and slow-relaxation components of NPQ (NPQF and NPQS, respectively) in 181 two varieties of rice, At-354 and Bg-352 at the early vegetative and flowering stages subjected to 0, 50 and 100 mM NaCl
Chapter 4
Table 4.1. PTOX and actin primers used in rt-PCR 214
Table 4.2. Proteins identified by the mass spectrometry. BN-PAGE separated protein 226 complexes of the thylakoid membranes isolated from T. salsuginea plants exposed to salt, drought and different irradiances
9 Abbreviations
A assimilation rate ADP adenosine diphosphate ATP adenosine triphosphate ATPase ATP synthase CET cyclic electron transport Car carotenoid Chl chlorophyll 3Chl* triplet excited state of chlorophyll
Ci internal CO2 concentration
Cyt b6f cyctochrome b6f complex Cyt f cyctochrome f ETR electron transport rate ETC electron transport chain Fd ferredoxin FQR ferredoxin-plastoquinone oxidoreductase FNR ferredoxin: NADP reductase Fo initial fluorescence level (obtained when QA is maximally oxidised) Fm maximum chlorophyll fluorescence level (obtained when QA is maximally reduced) Ft actual fluorescence intensity at any time FR far-red gs stomatal conductance k pseudo-first order rate constant for the reduction of oxidised P700 LED light emitter diode LHCI light harvesting complex I LHCII light harvesting complex II NADP oxidized nicotinamide adenine dinucleotide phosphate NADPH reduced nicotinamide adenine dinucleotide phosphate NPQ non-photochemical quenching P680 primary electron donor of PSII P680+ oxidised primary electron donor of PSII P700 primary electron donor of PSI
10 P700+ oxidised primary electron donor of PSI PC plastocyanin Pheo pheophytin PSI photosystem I PSII photosystem II PTOX plastid terminal oxidase PQ plastoquinone
PQH2 plastoquinol
QA primary quinone electron acceptor of PSII
QB secondary quinone electron acceptor of PSII qE high-energy-state quenching qI photoinhibitory quenching qP photochemical quenching qT state-transitions quenching RCs reaction centres ROS reactive oxygen species Rubisco ribulose 1,5-bisphosphate carboxylase/oxygenase RuBP ribulose bisphosphate SOD superoxide dismutase ΔpH pH gradient across the thylakoid membrane ΦPSII quantum yield of photosystem II
11 Abstract Regulation of Photosynthesis in plants under abiotic stress Sashila Sisimadhavi Abeyratne Abeykoon Walawwe A Thesis submitted to the University of Manchester for the degree of Doctor of Philosophy, 11 August 2014 Most plants complete their life cycle in a single location and therefore are affected by the changing environment. As a result, plants have evolved physiological and developmental adaptations to overcome stress. The work presented in this thesis has examined the regulation of photosynthetic electron transport in barley, rice and Thellungiella salsuginea. Barley is considered as a crop which is comparatively tolerant to soil salinity. The focus of this study was to evaluate the physiological responses of photosynthesis in barley under salinity and to characterize traits responsible for the regulation of photosynthesis. At low salt concentrations, barley plants protect PSII centres from excitation pressure by down-regulating the electron transport chain and maintaining ΔpH, by cyclic electron transport associated with PSI, to support non- photochemical quenching (NPQ). However, at the highest concentration of salt examined, this regulation starts to fail. The failure might result from a specific loss of PSI, resulting in reduced cyclic electron flow, or an increase in the leakiness of the thylakoid membranes, resulting in loss of ΔpH. The effects of salinity on the regulation of electron transport through Photosystem I and Photosystem II have been studied in two rice varieties from Sri Lanka. The regulation of photosynthesis in the salt-tolerant At-354 is more prominent than in the salt-sensitive Bg-352 when plants are exposed to salt. Exposure of Bg-352 to salt resulted in a substantial decrease in gas exchange, PSII photochemistry, leaf area and loss of chlorophylls. The decrease in the photosynthesis in AT-354 is caused by stomatal limitations, which restrict the CO2 entry into the plants, whereas the decrease of photosynthesis in Bg-352 is caused by non-stomatal limitations. Results suggest that At-354 protects PSII centres from excitation pressure by down-regulating the electron transport chain and maintaining ΔpH by cyclic electron transport associated with PSI to support NPQ. At high salt concentration, this regulation starts to fail in Bg-352. Tolerance to abiotic and biotic stress has evolved in many wild plant species, termed extremophiles. These plants contain essential genes which may used to improve crop production in changing environments. Thellungiella salsuginea is an extremophile, able to grow and reproduce in extreme environments. Stepien and Johnson (2009) identified a protein, known as the plastid terminal oxidase (PTOX) which acts as an alternative electron sink in T. salsuginea under salt stress. The current study showed that, in addition to salt, T. salsuginea showed increases in PTOX protein content and activity when exposed to drought, different growth irradiances and cold with high light. Semi-natural conditions also triggered the activity of PTOX. This study also showed that salt caused an up-regulation of PTOX gene transcripts in the leaves of salt treated T. salsuginea plants compared to control plants. Direct electron transport from PSII to PTOX and then to oxygen via the PQ pool accounted for up to 30% of total PSII electron flow in T. salsuginea (Stepien and Johnson, 2009). Efficient electron flow from PSII to PTOX would however, probably require co-location of these complexes in the same thylakoid fraction. To examine the location of PTOX in the thylakoid membrane, immunoblot analyses were performed, to test for changes in other protein complexes which may be associated with PTOX. In addition blue-native polyacrylamide gel electrophoresis and immunoblots were performed to isolate and detect the PTOX protein with any associated complexes. Although immunoblot analysis showed a prominent signal, mass spectrometry data did not allow identification of PTOX. This results suggests that further studies are needed to identify the precise localisation of the PTOX protein in the thylakoid membranes in T. salsuginea.
12 Declaration
I declare that no portion of the work referred to in the thesis has been submitted in support of an application for another degree or qualification of this or any other university or other institute of learning.
Copyright Statement i. The author of this thesis (including any appendices and/or schedules to this thesis) owns certain copyright or related rights in it (the “Copyright”) and s/he has given The University of Manchester certain rights to use such Copyright, including for administrative purposes. ii. Copies of this thesis, either in full or in extracts and whether in hard or electronic copy, may be made only in accordance with the Copyright, Designs and Patents Act 1988 (as amended) and regulations issued under it or, where appropriate, in accordance with licensing agreements which the University has from time to time. This page must form part of any such copies made. iii. The ownership of certain Copyright, patents, designs, trade marks and other intellectual property (the “Intellectual Property”) and any reproductions of copyright works in the thesis, for example graphs and tables (“Reproductions”), which may be described in this thesis, may not be owned by the author and may be owned by third parties. Such Intellectual Property and Reproductions cannot and must not be made available for use without the prior written permission of the owner(s) of the relevant Intellectual Property and/or Reproductions. iv. Further information on the conditions under which disclosure, publication and commercialisation of this thesis, the Copyright and any Intellectual Property and/or Reproductions described in it may take place is available in the University IP Policy (see http://www.campus.manchester.ac.uk / medialibrary / policies / intellectual-property. pdf), in any relevant Thesis restriction declarations deposited in the University Library, The University Library’s regulations (see http://www. manchester. ac. uk / library / aboutus / regulations) and in The University’s policy on presentation of Theses
13 Acknowledgements
First and foremost my gratitude goes to my supervisor, Dr Giles Johnson. This thesis was made possible due to the masterly guidance of him. I am thankful to him for the patient guidance, encouragement and advice he has provided throughout my PhD. I am fortunate to have Dr Patrick Gallois as my advisor, who cared so much about my work and who responded to my questions and queries so promptly.
Many thanks to the University of Manchester for giving me this opportunity to do the PhD and for the funding. I would like to thank FFWG for providing me funds during my third year. I am thankful to Rice Research Institute in Sri Lanka for providing me rice seeds and Mr Kulunusen Yasakethu, former director of the Agriculture Research Centre at Ganoruwa, Sri Lanka and my beloved uncle for helping me to get rice seeds from Sri Lanka and all his efforts during the quarantine procedure. I specifically owe thanks to Furzani, Beth, Matt and Chuks for their enormous support. A special thank goes to friends, Xun and Yaomin, who helped me throughout my molecular biological experiments and giving me moral support. Many thanks to all lab members in D.3503 for our lovely stress busting lunch-time discussions.
To my lovely husband, Isuru, thank you so much for your support, understanding and love during these most stressful and the most fulfilling four years of my life. Thank you for being there for me and being the best husband in the world. I love you.
Family is the most important thing in my life. So I would like to thank my dad, who sacrificed his own 'PhD dream' to fulfil my dream and providing support mentally and financially. I love you so much and missed you terribly. My loving mum, for her unconditional love and support throughout my life. A special thank goes to my one and only brother, who gives me enormous support and encouragement. Last but not least many thanks to my lovely in-laws, who are always willing to help when ever I needed.
Dedicated to the memory of my beloved father Colin Abeyratne
14 Chapter 01
General Introduction
15 1.1. Introduction
The effects of abiotic stress, due to changes in the physico-chemical environment, are reflected at quantitative and qualitative levels in all agricultural lands worldwide (Boyer, 1982). Although many stress conditions, such as drought, heat, salinity and low temperatures have been the subject of intense research, in the field, crops are always subjected to combinations of different abiotic stresses
(Mittler, 2006). Plants show complex responses to stress. The responses of plants to combinations of stresses can be different to those seen when each stress applied individually (Mittler, 2006;
Cramer et al., 2011).
Photosynthesis is the most important physiological function of a plant which has a direct effect on the plant growth and is highly susceptible to environmental stress (Chaves et al., 2003; Flexas et al.,
2004; Chaves et al., 2009; Lawlor and Tezara, 2009; Pinheiro and Chaves, 2011). Abiotic stress including salt and drought suppresses photosynthesis by affecting photosynthetic pigments, soluble proteins, proteins in thylakoid membranes, the electron transport chain, photophosphorylation and
CO2 fixation. Inhibition of photosynthesis disrupts plant growth (Sudhir and Murthy, 2004).
However, depending on their tolerance level, plants show different responses to abiotic stress.
Therefore, it is important to study the effects of abiotic stresses on plants to understand the physiology of stress responses in plants.
This thesis focuses on analysing the effects of abiotic stress on photosynthesis of plants and the regulation of photosynthesis under abiotic stress. Chapter 1 provides a literature review of topics related to the study. As barley is a relatively salt tolerant crop (Munns and Tester, 2008), a study of barley provides insight into the regulatory mechanisms of photosynthesis. Chapter 2 discusses the regulation of photosynthesis in barley under salt stress. Chapter 3 focuses on a physiological evaluation of salinity stress in two rice varieties from Sri Lanka. Sri Lanka is a rice growing country
16 and subjected to crop losses every year due to soil salinity. Therefore, the physiological evaluation of photosynthesis of two extensively use rice varieties under salinity gives a better understanding of the salt tolerant and sensitive traits of rice which can be implemented for the future development of rice cultivars.
Some plants have different regulatory mechanisms to overcome photodamage. Plants like
Thellungiella salsuginea (T. salsuginea) show the presence of an alternative electron sink, the plastid terminal oxidase (PTOX), under salt stress (Stepien and Johnson, 2009). Chapter 4 focuses on detecting PTOX protein using western blot analysis, examining the transcriptional regulation using rt-PCR and analysing the activity under ambient and low oxygen concentrations when T. salsuginea was challenged with abiotic stresses, including drought, salt, different growth light and cold with high light. PTOX is assumed to interact with the PQ pool independently. However, there is a possibility that it may be a subunit of some larger thylakoid protein complex and that the association may be depend on the environmental conditions (McDonald et al., 2011). The study also examines the effects of abiotic stresses on the protein content of the other photosynthetic complexes, such as PSII and cytochrome b6f, which are predicted to be associated with PTOX. Blue native gel electrophoresis and mass spectrometry analysis were used to examine the possible location of PTOX and other associated complexes in the thylakoid membrane.
17 1.2. Photosynthesis
Photosynthesis is the process converting light energy to chemical energy and storing the energy in the bonds of sugars and other organic compounds. Carbohydrates are synthesized from water and carbon dioxide and oxygen is released as a by-product. In plants and algae, the process of photosynthesis occurs in specialized organelles known as chloroplasts which contain the photosynthetic apparatus. Photosynthesis mainly occurs in mesophyll cells in plant leaves.
Photosynthesis is one of the major metabolic processes which is sensitive to environmental stresses.
It is important to regulate the energy flow to optimise carbon fixation and prevent light induced damage. Photosynthesis consists of two major processes: the light-dependent electron transport chain and the light-independent carbon fixation cycle or the Calvin-Benson-Bassham cycle.
1.2.1. Light capture and Electron Transport Chain
The light reactions take place in the thylakoid membranes, inside the chloroplast. Light energy is captured by a series of pigments localized in light harvesting complexes (LHC) and reaction centers
(RC) in two complexes: photosystem I (PSI) and photosystem II (PSII) (Figure 1.1). Plants absorb light mainly by chlorophylls. In addition, plants also contain other light harvesting pigments, carotenes and xanthophylls. Most of the PSII centers are localized in appressed membrane regions, called the grana stack of the thylakoid membranes. In contrast, PSI centers are mainly localized in the non-appressed regions (margin of the grana stacks) (Albertsson, 1995; Albertsson, 2001) or in stroma lamellae (Kirchhoff et al., 2000). Light energy, captured by pigments in LHC, is transferred to reaction centers (RCs) through resonance energy transfer. In the RCs, the absorbed energy causes transition in specialized chlorophyll-a molecules, P680 (in PSII) and P700 (in PSI), from a ground state to an excited state, where an electron is promoted to an orbital with greater potential energy.
18 This electron can then be transferred to another molecule, through charge separation. Donation of this electron from excited P680 to nearby pheophytin (Pheo) can be thought of as the first step in electron transport chain (Klimov et al., 1977; Klimov et al., 1979; Groot et al., 2005; Holzwarth et al., 2006). PSII forms a dimeric supercomplex and has a molecular weight of 1400 kDa (Caffarri et al., 2009). The monomer contains about 40 different proteins most of which are permanent parts of the structure and others expressed or associated with it during stress conditions or assembly and degradation (Shi et al., 2012).
PSII is a multi subunit pigment-protein complex with two moieties: the core, highly conserved in all photosynthetic organisms, contains the major cofactors of electron transport and the outer light harvesting antenna complex consists of most of the light absorbing pigments and provides the core with excitation energy (Nield et al., 2000; Büchel and Kühlbrandt, 2005; Dekker and Boekema,
2005; Croce and van Amerongen, 2011; Umena et al., 2011) (Figure 1.2). In plants and eukaryotic algae the light harvesting complex (LHC) proteins which are encoded by a multigenic family are located in the thylakoid membrane (Jansson, 1999). LHC proteins can be categorized as LHCI (in
PSI) and LHCII (in PSII) which encoded by Lhca or Lhcb genes, respectively. Among two types of isoforms of LHCII, monomers, CP29 (Lhcb4), CP26 (Lhcb5) and CP24 (Lhcb6) are less abundant, located close to PSII core complex and have different pigment composition (Croce et al., 2002;
Dekker and Boekema, 2005; Passarini et al., 2009; Pan et al., 2011). The most abundant LHCII consists of 3 gene products (Lhcb1, Lhcb2 and Lhcb3) which are organized as heterotrimers (Ben-
Shem et al., 2003; Liu et al., 2004; Standfuss et al., 2005; Drop et al., 2014). The most abundant pigments found in LHCII are chlorophyll-a and chlorophyll-b, which are involved in the energy capture and transfer toward P680 (Barber and Archer, 2001). In addition to these, pigments including lutein, neoxanthin and the xanthophyll cycle carotenoids, violaxanthin, antheraxanthin and zeaxanthin are involved in energy transfer, energy dissipation and the scavenging of reactive
19 oxygen species (ROS) (Niyogi et al., 1997; Croce et al., 1999; Ballottari et al., 2012; Grewe et al.,
2014).
Figure 1.1. Schematic model of the major protein complexes involved in electron transport chain in
photosynthesis (retrieved from http://macromol.sbcs.qmul.ac.uk/showcase/showcase.html.)
PSII consists of the core reaction centre (RCII) where the charge separation take place and the outer
light harvesting complex (LHCII) where the majority of solar energy is captured and transferred to
the core. The multi-nuclear Mn4Ca cluster, the water splitting complex or the oxygen-evolving
complex (OEC) is responsible for oxidizing water to molecular oxygen. Cytochrome b6f (Cyt b6f)
complex mediates the electron transport between PSII and PSI which is coupled with proton
transfer from the stroma to the lumen to generate a pH gradient across the membrane. Similar to
PSII, PSI consists of two membrane complexes as the reaction centre (RCI) core and the outer light
harvesting complex (LHCI). RCI is where most of the light capturing and the charge separation take
place and LHCI acts as an additional antenna system that maximizes light harvesting and transport
the energy to the core complex. The chloroplast ATP synthase consists of two parts, Fo rotor
embedded in the membrane and the catalytic F1 extended towards the stroma. ATP synthase
generates ATP through translocating proton across Fo domain.
20 The PSII core, which consists of the complex of the reaction center with D1, D2 and cytochrome b-
559 (Cyt b-559) subunits, generates the redox potential necessary to drive water splitting (Dau et al., 2012). Other than the reaction center, the core also contains the chlorophyll (Chl) a-binding antenna complexes, CP43 and CP47 (Dekker and Boekema, 2005; Dang et al., 2008). D1 and D2 forms a quasi-symmetrical complex with several cofactors (Ishikita and Knapp, 2006; Saito et al.,
2013). Several extrinsic proteins, including PsbO, PsbQ, PsbP and PsbR (in plants and algae) are associated with the lumenal side the core complex (Ifuku et al., 2011; Dau et al., 2012). These proteins form the oxygen-evolving complex (OEC) or water splitting complex (Ifuku et al., 2011;
Dau et al., 2012). In a PSII monomer, apart from the protein subunits, 35 chlorophylls, two pheophytins, 11 β-carotenes, more than 20 lipids, two plastoquinones, two haem irons, one non- haem iron ions, 4 manganese atoms, 3-4 calcium atoms (one of which is in the Mn4Ca cluster), 3
- chlorine ions (Cl ) (two of which are in the vicinity of the Mn4Ca cluster) and one bicarbonate ion have been identified (Umena et al., 2011). There are two redox active tyrosine residues in PSII as
D1-Tyr161 (tyrosine Z, TyrZ) and D2-Tyr160 (tyrosine D, TyrD) both of which can provide electrons to P680+. (Barry and Babcock, 1987; Debus et al., 1988; Vermass et al., 1988). However,
TyrZ, which has D1-His190 as an H-bond partner, is more kinetically competent than TyrD which has D2-His189 as H-bond partner, and mediates proton-coupled electron transfer from Mn4CaO5 to
P680+. (Ishikita and Knapp, 2006; Saito et al., 2013). Cyt b-559 is a ubiquitous component of PSII and located close to D1 and D2 subunits of PSII (Nanba and Satoh, 1987). Cyt b-559 consists of heterodimer of α (PsbE) and β (PsbF) subunits with the haem molecule joined together by single histidine residue in each subunit. According to the crystal structure of cyanobacterial PSII, this component is located next to the D2, where the haem molecule is positioned towards the cytoplasm.
The approximate distance between Cyt b-559 and the binding site of QB is 25 Å and the Cyt b-559 is 11.6 Å from a carotenoid CarD2 (bound to the D2 subunit) (Ferreira et al., 2004; Loll et al., 2005;
Guskov et al., 2009; Umena et al., 2011). Although many studies have been performed to find the
21 physiological role of Cyt b-559, it still remains unresolved. A recent study on the His-H23C, a mutant of Chlamydomonas reinhardtii (C. reinhardtii) (His ligand to the haem of PsbE subunit is replaced by a Cys residue) suggested that Cyt b-559, plays a major role in the assembly, repair and maintenance of the complex and improves the electron transport on the acceptor side of PSII
(Hamilton et al., 2014). The Mn4Ca cluster in the OEC is responsible for the oxidizing water to molecular oxygen. The mechanism of water splitting in this cluster occurs through four electron oxidation process with five intermediate 'S' states, 4 of these are meta-stable (S0, S1, S2, S3) and one short-lived state (S4) with higher mean oxidation levels (Satoh et al., 2005; Jin et al., 2014).
Apart from four manganese atoms and single calcium atom, OEC consists of five oxygen atoms
(Umena et al., 2011). The X-ray structure of PSII published by Umena et al. (2011) clearly showed the positions of the manganese and calcium atoms in the OEC, the oxo-bridges connecting the metal atoms and a number of coordinated water molecules.
In PSII charge separation, one molecule of chlorophyll absorbs one photon and loses one electron to
Pheo, which then transfers to plastoquinone A (QA), the primary accepting quinone at the stromal side of the protein. The charge separate states of PSII RC contains four molecules of chlorophylls
(ChlD1, ChlD2, PD1, PD2) and two molecules of Pheo (PheD1, PheD2) (Zouni et al., 2001; Ferreira et al., 2004; Loll et al., 2005; Guskov et al., 2009). A study by Romero et al. (2010) showed a presence of two different charge separation pathways in PSII. They suggested that, at least two different excited states, (ChlD1PheD1)* and (PD1PD2ChlD1)* initiate these pathways in PSII RC. From
QA, the electron is transferred to a second plastoquinone molecule in the QB site of PSII (Klimov et al., 1978). The intermediate QB semiquinone, which is formed is stable in the QB site for several seconds (Mitchell, 1979; Diner et al., 1991; Schuurmans et al., 2014). Following a second charge separation, reduction of the semiquinone results in formation of a reduced plastoquinol (PQH2) which accepts protons from the stroma of the chloroplast.
22 Figure 1.2. Schematic model representing the proteins of light harvesting complex II (LHCII) and reaction centre core (RCII) of photosystem II (PSII) (retrieved from http://macromol.sbcs. qmul.ac.uk/oldsite/psIIimages/PSII.html). PSII is a multi subunit pigment-protein complex with two moieties: the core, highly conserved in all photosynthetic organisms, contains the major cofactors of electron transport and the outer light harvesting antenna complex consists of most of the light absorbing pigments and provides the core with excitation energy. Lhcb1, Lhcb2 and
Lhcb3 are organized as heterotrimers and Lhcb4, Lhcb5 and Lhcb6 are organized as monomers.
The PSII core consists of the complex of the reaction center with D1, D2 and cytochrome b-559
(Cyt b-559) subunits. Other than the reaction center, the core also contains the chlorophyll (Chl) a-binding antenna complexes, CP43 and CP47. Several extrinsic proteins, including PsbO, PsbQ,
PsbP and PsbR (in plants and algae) are associated with the lumenal side the core complex (Ifuku et al., 2010; Dau et al., 2012). These proteins form the oxygen-evolving complex (OEC) or water splitting complex (Ifuku et al., 2011; Dau et al., 2012)
23 Following each charge separation P680 is re-reduced by an electron from a tyrosine molecule
(TyrZ/YZ) which is in turn reduced by water. Water photolysis takes place in the oxygen evolving or water splitting complex associated with PSII. This process releases oxygen and protons (H+) into the thylakoid lumen. The mobile pool of plastoquinol acts as an electron mediator between PSII and cytochrome b6f complex (Cyt b6f) (Albertsson, 1995; Kirchhoff et al., 2000; Albertsson, 2001).
Cytochrome b6f (Cyt b6f, plastohydroquinone:plastocyanin oxidoreductase) (molecular weight of
220 kDa) is partially homologous to the cytochrome bc1 found in mitochondria. Cyt b6f mediates electron transport between PQH2 and plastocyanin (PC) which is coupled with proton transfer from the electrochemically negative (n) to the positive (p) side of the complex which generates a pH gradient across the thylakoid membrane (Baniulis et al., 2009). This complex consists of 8 subunits,
13 transmembrane helices and 7 prosthetic groups (4 haems, 1 (2Fe-2S) cluster, 1 chlorophyll-a and
1 carotene) (Alric et al., 2005; Yamashita et al., 2007). Four large subunits (18-32 kDa), cytochrome f (Cyt f), cytochrome b6 (Cyt b6), the Rieske iron-sulphur protein (2Fe-2S protein) and subunit IV and four small hydrophobic subunits, PetG, PetL, PetM and PetN form the functional dimer of Cyt b6f (Mitchell, 1975; Croft and Wraight, 1983; Whitelegge et al., 2002). Among these, Cyt f and Cyt b6 contain haem prosthetic groups. Haem f (c-type haem) is covalently attached to the Cyt f on the lumenal face (positive side) and haem ci/cn is covalently attached to the Cyt b6 polypeptide on the negative side of the complex (the quinone binding site, Qi) (Kurisu et al., 2003; Stroebel et al.,
2003). The bp and bn, b-type haems (bis-histidine ligated haems) are found on the p and n-sides of the Cyt b6, respectively (Hasan et al., 2013). These two haem molecules are represented as bH (H for high mid point potential) and bL (L for low mid point potential) in Stroebel et al. (2003). Figure
1.3.a showed the electron and proton transport pathway in Cyt b6f (Q cycle) described by Stroebel et al. (2003) and Figure 1.3.b by Baniulis et al. (2013).
24 a
e- b 2H+ Fd FNR e- Stroma (n)
PQH2 PQ bn /cn Low potential chain e-
bp e- PQH2 PQH- PQH PQ-.
Lumen (p)
H+ e- H+
e- e- Rieske Fe-S protein Cyt f PC High potential chain
Figure 1.3. The model of Q cycle representing the electron and proton transport in the Cyt b6f (a) by
Stroebel et al. (2003) (retrieved from http://macromol.sbcs.qmul.ac.uk/oldsite/psIIimages/cytb6f
.html) (b) by Baniulis et al. (2013). One electron is transferred from plastoquinol (Qo site) (PQH2) the Rieske iron-sulphur protein and cytochrome f (Cyt f) on the electropositive side (p-side) of the membrane. This is a high potential chain. Two protons are released to the aqueous lumen phase. The second electron from PQH2 transport through two b-type haems, bp and bn, and cn. Electrons are input into Cyt b6f through ferredoxin or perhaps through FNR. Uptake of protons from the stroma generate a pH gradient across the membrane. 25 Electrons are then transferred to PC, a copper containing mobile protein molecule in the thylakoid lumen, via Cyt b6f. The lateral flow of electrons between the appressed region (Cyt b6f) and the non-appressed region (PSI) is mediated by PC (Kirchhoff et al., 2000). Protons (H+) from plastoquinol released into the lumen, build a pH gradient across the thylakoid membrane. As in
PSII, a specialized chlorophyll-a, P700 in PSI, absorbs light energy and causes charge separation in the reaction centers. Electrons are then transferred to the iron-sulphur protein ferredoxin (Fd) via iron-sulphur centers in PSI, then to ferredoxin-NADP+ reductase (FNR) and used to reduce NADP+ to NADPH. Movement of H+ ions from the lumen to the stroma is used by ATP synthase to drive the production of ATP. The products of the electron transport chain, NADPH and ATP are fed into the Calvin-Benson cycle.
PSI is a monomer in plants (Scheller et al., 2001; Ben-Shem et al., 2003) and a trimer in cyanobacteria (Boekema et al., 1987; Fromme and Witt, 1998) with two membrane complexes as the reaction centre (RC) core and the outer light harvesting complex (LHC) (Figure 1.4). According to Nelson and Yocum (2006), the photochemical quantum yield of PSI is close to 1.0. For this reason, PSI is considered as the most efficient light capturing and energy converting device in nature (Amunts and Nelson, 2009). Amunts and Nelson (2009) identified that a PSI-LHCI supercomplex consists of 13 proteins, where four of them are the peripheral LHC proteins (Lhca1,
Lhca2, Lhca3, Lhca4), 45 transmembrane helices, 3 stroma-exposed subunits, 1 lumenal subunit,
168 chlorophylls. Apart from that, 3 Fe4-S4 clusters, 2 phylloquinones, and 5 carotenoids.
Additional LHC proteins, Lhca5 and Lhca6 were identified in Arabidopsis thaliana (Jansson, 1999).
A study by Peng and Shikanai (2011) suggested that these additional LHC proteins are involved in
NAD(P)H dehydrogenase (NDH)-PSI supercomplex formation promoting cyclic electron flow. The reaction centre is located in the core of the complex and consists of a heterodimer of the two large transmembrane protein subunits, PsaA and PsaB, comprising 22 transmembrane helices which
26 provide binding sites for the donor P700, and the acceptors Ao, A1 and Fx (Amunts and Nelson,
2009).
The RC is where most of the light capturing and the charge separation take place and LHCI acts as an additional antenna system that maximizes light harvesting and transfer the energy to the core complex (Chitnis, 2001; Amunts and Nelson, 2009). PsaA and PsaB associated with the three stromal subunits PsaC, PsaD, and PsaE which provide a binding groove for ferredoxin (Fd)
(Amunts and Nelson, 2009). Flash absorption spectroscopy study performed by Fischer et al.
(1999) identify two different kinetic phases in the reduction of soluble ferredoxin by PSI. Excited
P700 transferred electrons to Fd through a chlorophyll molecule Ao, a phylloquinone A1 and three terminal Fe4-S4 clusters, Fx, FA and FB (Rutherfold and Mullet, 1981; Sétif et al., 1981; Fromme and Mathis, 2004). FNR mediates the electron transport between Fd (one electron carrier) and
NADP+ (two electron carrier) at the end of the electron transport (Equation 1) (Hurley et al., 2002;
Musumeci et al., 2012). In addition, FNR also involves in the electron transport in isoprenoid biosynthesis, nitrogen fixation, steroid metabolism, xenobiotic detoxification, oxidative-stress response and iron-sulfur cluster biogenesis (Carrillo and Ceccarelli, 2003; Ceccarelli et al., 2004;
Medina and Gomez-Moreno, 2004; Röhrich et al., 2005; Seeber et al., 2005).
Equation 1:
+ + 2Fdreduced + NADP + H 2Fdoxidised + NADPH
27 Figure 1.4. Schematic model representing the proteins of light harvesting complex I (LHCI) and reaction centre core (RCI) of photosystem I (PSI) (retrieved from http://macromol.sbcs. qmul.ac.uk/oldsite/psIIimages/PSI.html). PSI-LHCI supercomplex consists of 13 proteins which four of them are the peripheral LHC proteins (Lhca1, Lhca2, Lhca3, Lhca4), 45 transmembrane helices, 3 stroma-exposed subunits, 1 lumenal subunit, 168 chlorophylls. Apart from that, 3 Fe4-
S4 clusters, 2 phylloquinones, and 5 carotenoids (Amunts and Nelson, 2009). Lhca5 and Lhca6 are involved in NAD(P)H dehydrogenase (NDH)-PSI supercomplex formation promoting cyclic electron flow (Peng and Shikanai, 2011). PsaA and PsaB form a heterodimer in the reaction centre core and provide a binding site for the donor P700 and the acceptors Ao, A1 and Fx. PsaA and
PsaB associated with subunits PsaC, PsaD, and PsaE which provide a binding groove for ferredoxin (Fd) (Amunts and Nelson, 2009).
28 The major complexes including PSII, PSI and Cyt b6f are involved in generating a pH gradient across the thylakoid membrane through light-driven charge separation (Nelson and Ben-Shem,
2004). The chloroplast ATP synthase (chloroplast F1Fo synthase, cF1Fo) generates ATP by using this proton gradient, with protons being translocated across the Fo sector (Daum et al., 2010)
(Figure 1.5). According to low resolution electron microscopy analysis, the structure of cF1Fo is similar to the homologous bacterial and mitochondrial enzymes, and has an Fo rotor embedded in the membrane and a catalytic F1 extended towards the stroma (Stock et al., 1999; Mellwig and
Böttcher, 2003). The catalytic F1 which is too large to fit in the stromal gap between stacked grana thylakoids restricts the distribution of cF1Fo only into the nonstacked regions of the thylakoid membranes (Oleszko and Moudrianakis, 1974; Miller and Staehelin, 1976; Staehelin et al., 1976;
Mellwig and Böttcher, 2003). The F1 domain is made up of subunits, α3, β3, γ, δ and ε (Boekema et al., 1998). Fo, the membrane embedded domain, consists of subunits a, b, b', and a c-ring rotor, which are also known as IV, I, II, III, respectively (Seelert et al., 2000; Varco-Merth et al., 2008).
The α3β3 complex is a hexamer with a central cavity which allows the penetration of the γ rotor shaft (Abrahams et al., 1994). γ and ε forms the rotor shaft which is called the central shaft (Junge et al., 1997). Rotation of the γ subunit within the α3β3 cavity causes conformational changes in the three catalytic sites at the α-β interfaces. This drives the ATP production (Gibbons et al., 2000).
There are two stalks attached to F1 and Fo domain which help them to held together. One is the central rotating shaft consists of subunits, γ and ε and the other is a thin stalk made up of b, b' and δ
(Junge et al., 1997). Subunit c forms a ring structure which interacts with foot of central stalk subunits (Stock et al., 1999). a and c subunits associated together for the proton pumping in the ATP synthase the number of copies of these a and c (varies from 8-15) depends on the species (Stock et al., 1999; Watt et al., 2010). A recent study on the crystallographic structure of the c-ring from spinach chloroplast showed that each c-ring contains 14 monomers in the asymmetrical unit
(Balakrishna et al., 2014).
29 Figure 1.5. Schematic model representing the chloroplast ATP synthase (retrieved from http://macromol.sbcs.qmul.ac.uk/oldsite/psIIimages/atpase.html). It has a Fo rotor embedded in the membrane and a catalytic F1 extended towards the stroma. The F1 domain is made up of subunits, α3, β3, γ, δ and ε (Boekema et al., 1998). Fo, the membrane embedded domain, consists of subunits a, b, b', and a c-ring rotor, which are also known as IV, I, II, III, respectively (Seelert et al., 2000; Varco-Merth, 2008). Rotation of the γ subunit within the α3β3 cavity causes conformational changes in the three catalytic sites at the α-β interfaces. This drives the ATP production (Gibbons et al., 2000). Subunit c forms a ring structure which interacts with foot of central stalk subunits (Stock et al., 1999) a and c subunits associated together for the proton pumping in the ATP synthase. OSCP is the ATP synthase delta subunit, which is a part of the stalk that holds the F1 complex catalytic core (Davies et al., 2012).
30 The reducing power build via linear electron transport is also used to reduce thioredoxin (TRX), which leads to activate several enzymes in the Calvin-Benson cycle (Rochaix, 2013). Apart from acting as a light energy collector and converter, the photosynthetic apparatus also plays a vital role as a sensor which regulates the electron transport chain according to the changes in its environment, such as light quality and intensity, water availability, temperature and metabolic needs (Rochaix,
2013). These changes affect the redox poise of the PQ pool and the Fd/TRX pool and the pH gradient across the thylakoid membranes, which is sensed by the photosynthetic machinery
(Rochaix, 2013). This leads the photosynthetic apparatus to trigger regulatory processes to rebuild the optimal redox poise in the electron transport chain.
1.2.2. Cyclic Electron Transport
In addition to the linear electron transport chain, cyclic electron transport (CET) is found in photosynthetic organisms including plants, algae and cyanobacteria (Fork and Herbert, 1993;
Bendall and Manasse, 1995; Heber, 2002; Allen, 2003). This electron transport pathway was first identified by Arnon and co-workers (1954) as cyclic phosphorylation. CET helps to build a pH gradient across the thylakoid membranes (Heber and Walker, 1992; Munekage et al, 2004; Shikanai,
2014). Both PSII and PSI are involved in linear electron transport to produce NADPH and ATP, whereas CET is driven by PSI alone and produces ATP (Shikanai, 2007; Shikanai, 2014). Due to the nature of cyclic processes, showing no net flux, CET has been difficult to study, especially in leaves
(Johnson, 2005). There are suggested to be two major functions of CET. (i) ATP synthesis: CET generates a pH gradient which supports production of ATP to, overcome imbalances of the ATP:
NADPH (Seelert, 2000; Allen, 2003; Joliot et al., 2006; Joliot and Johnson, 2011). (ii) CET encourages an increase of non-photochemical quenching (NPQ) by increasing the ΔpH across the thylakoid membranes and inhibits the production of reactive oxygen species (ROS) (Heber and
31 Walker, 1992; Clarke and Johnson, 2001; Golding and Johnson, 2003; Joliot and Johnson, 2011).
There are two commonly recognised pathways of CET (Figure 1.6) (Munekage et al., 2004). In one, ferredoxin (Fd), which is the terminal electron acceptor of the ETC, reduces plastoquinone in the presence of a putative enzyme ferredoxin quinone reductase (FQR) and electrons are fed into Cyt b6f complex (Johnson, 2005). PC, Cyt b6f complex, PSI and Fd are involve in this pathway.
However, the exact pathway is yet to be defined (Bendall and Manasse, 1995; Johnson, 2011; Hertle et al, 2013). This pathway is sensitive to the electron transport inhibitor antimycin A (AA) and referred to as AA-sensitive cyclic electron flow (Bendall and Manasse, 1995; Hertle et al, 2013;
Shikanai, 2014). In mitochondria, AA binds to and blocks the quinone binding site (QN) on the cytochrome bc1 complex. However, this has not observed in chloroplast (reviewed in Bendall and
Manasse, 1995; Shikanai, 2014). The presence of an AA-sensitive putative enzyme FQR was postulated by Moss and Bendall 30 years ago (Moss and Bendall, 1984), but it has not so far been identified conclusively through biochemical or genetic approaches (Joliot and Johnson, 2011; Hertle et al, 2013).
A study by Munekage et al. (2002) showed PGR5 (proton gradient regulation 5), a small thylakoid protein, is involved in the Fd-dependent CET and supports NPQ by maintaining a ΔpH in the thylakoid membranes. Similarly, another intrinsic membrane peptide, PGRL1 (proton gradient regulation 5 like 1) was identified in Arabidopsis thaliana as involved in this process (DalCorso et al., 2008; Iwai et al., 2010; Joliot and Johnson, 2011). A study by Joliot and Johnson (2011) showed that plants with reduced level of FNR and plants lacking PGR5 have a reduced CET. However, the exact role of these two proteins in AA-sensitive CET is yet to identified (Hald et al., 2008; Nandha et al., 2007; Suorsa et al., 2012). A study by Sugimoto et al. (2013) found that altering a single amino acid in PGR5 in Arabidopsis thaliana confers resistance to antimycin A in CET around PSI
32 in leaves and ruptured chloroplasts. These findings suggested that the function of PGR5 or a protein closely associated with PGR5 (probably PGRL1) is inhibited by antimycin A (Sugimoto et al, 2013;
Shikanai, 2014). Consistent with these findings, work from Hertle et al. (2013) showed that electron transport from recombinant PGRL1 to DMBQ (quinone 2, 6-dimethyl-p-benzoquinone, which is a
PQ analog) was inhibited by antimycin A.
Hertle and co-workers (2013) proposed that in higher plants, PGRL1 is the putative enzyme FQR.
According to early studies on FQR, it should be sensitive to antimycin A (Bendall and Manasse,
1995), loosely associated with PSI (Bendall and Manasse, 1995), contain redox active moieties
(Bendall and Manasse, 1995; Shikanai, 2007), be present in relatively high concentrations (Mills et al., 1978; Joliot and Johnson, 2011) and be responsible for accepting electrons from Fd and donating them to PQ (Moss and Bendall, 1984). Consistent with these, Hertle et al. (2013) showed that antimycin A inhibits PGRL1 oxidation, both PGRL1 and PGR5 should be associated with PSI because it was found in PSI preparation (DalCorso et al., 2008), PGRL1 has redox active six cysteine (Cys) residues which are essential for PGRL1 heterodimerization, homodimerization, iron binding and the formation of disulfide bridges (Petroutsos et al., 2009) and PGRL1 is found in relatively high concentrations. In addition, this study proposed that PGRL1 is capable of mediating electron transport from Fd to PQ. PGR5 is suggested to be involved in the transfer of electrons from
Fd to PGRL1 in Arabidopsis thaliana (DalCorso et al., 2008; Hertle et al., 2013). PGRL1 interacts physically with PGR5 and forms heterodimers or, in the absence of PGR5 forms stable homodimers
(DalCorso et al., 2008; Hertle et al., 2013). However, PGRL1 is more abundant than PGR5 implying that multiple PGRL1 molecules are associated with one PGR5 (Hertle et al., 2013).
PGRL1 is mostly located in the appressed regions of thylakoids (DalCorso et al., 2008; Iwai et al.,
2010). Therefore, according to the proposed current model of antimycin A-sensitive CET in plants, electron transfer from Fd to PQ is mediated by a PGRL1-PGR5 complex and electrons are then
33 transferred to the Cyt b6f complex (Figure 1.6.a) (Hertle et al., 2013; Shikanai, 2014). However, apart from the involvement of PGRL1 in CET, it is also interacts with plastidic type I signal peptidase 1 (Plsp1), an isoform of the thylakoid processing peptidase (TPP), implying that PGRL1 may also be involved in a divergent function (Endow and Inoue, 2013). Therefore, the mechanism and the regulatory process of CET is still not fully understood (Shikanai, 2014).
+ A PSI-LHCI-LHCII-Fd-NADP reductase (FNR)-Cyt b6f-PGRL1 supercomplex mediating CET has been observed in green alga, C. reinhardtii (Iwai et al., 2010). Formation of this supercomplex depends on state transitions, where the LHC of the two photosystems are remodelled according to changes in light conditions which lead to (de)phosphorylation of LHCII (Finazzi et al., 2002; Iwai et al., 2010; Rochaix, 2013). PGRL1 present in C. reinhardtii does not act as the FQR but acts to stabilize this supercomplex and provides an interface between PSI and Cyt b6f (Figure 1.6.b) (Iwai et al., 2010; Hertle et al., 2013). However, the presence of such complex in higher plants has not been observed (Wollman and Bultẽ, 1989; Breyton et al., 2006; Iwai et al., 2010). Most importantly,
PGR5 is not found in CET in C. reinhardtii, but is an essential component of CET in plants
(DalCorso et al., 2008; Iwai et al., 2010). This raised the question that how electrons from Fd are transported to PQ without the involvement of PGR5 (Shikanai, 2014). CET in C. reinhardtii is also insensitive to antimycin A, suggesting it may be mechanistically different (Iwai et al., 2010; Hertle et al., 2013). Apart from PSI, LHCI, LHCII, FNR, Cyt b6f, and PGRL1, the supercomplex mediating CET in C. reinhardtii contains a chloroplast localized Ca2+ sensor and anaerobic response
1 (ANR1) (Terashima et al., 2012). Presence of these components associated with the CET supercomplex, suggests the activation of electron transport via Ca2+ signalling (Terashima et al.,
2012). A recent study by Dang et al. (2014) showed that both mitochondrial respiration and the
Mehler reaction supplies extra ATP for photosynthesis in the pgrl1 mutant of C. reinhardtii which is deficient in PGRL1 mediated CET.
34 FNR is a flavoprotein (molecular weight of approximately 35kDa), which are hydrophilic proteins with two structural domains, each having approximately 150 amino acids (Arakaki et al., 1997;
Kurisu et al., 2001; Aliverti et al., 2008; Paladini et al., 2009, Mulo, 2011). The flavin adenine dinucleotide (FAD) group binds to the amino-terminal domain which is an anti-parallel β-barrel
(Kurisu et al., 2001; Paladini et al., 2009). This concave region is the Fd recognition site (Kurisu et al., 2001). FAD is a non-covalently bound cofactor which acts as a redox centre (Paladini et al.,
2009). The carboxyl-terminal domain is the NADP+ site with a characteristic α-helix-β strand fold
(Paladini et al., 2009). FNR forms an electron transfer complex with Fd where the two prosthetic groups, the 2Fe-2S cluster in Fd and the FAD in FNR are sufficiently close for direct electron transfer from Fd to FNR (Kurisu et al., 2001). FNR is suggested to be exists either as a soluble pool in the chloroplast stroma or bound to the inner envelope membrane of the chloroplast via a subunit of chloroplast protein import machinery, Tic62 (Böhme, 1977; Fredricks and Gehl, 1982; Matthijs et al., 1986; Küchler et al., 2002; Balsera et al., 2007; Stengel et al., 2008). The precise location of
FNR in the photosynthetic apparatus is still controversial. Early studies suggested that FNR is associated with the thylakoid membrane through a base protein of molecular weight of approximately 16.5-17.5 kDa (Vallejos et al., 1984; Ceccarelli et al., 1985; Chan et al., 1987).
Studies performed by Clark et al. (1984) and Zhang et al. (2001) showed the association of FNR with Cyt b6f in Fd-dependent cyclic electron transport. Andersen et al. (1992) showed a possible association of FNR with the PsaE subunit of PSI. Studies of Guedeney et al. (1996) and José Quiles and Cuello (1998) showed FNR is associated with the NDH complex. Several studies have shown the co-localization of FNR with a Calvin cycle enzyme, glyceraldehyde-3-phosphate dehydrogenase
(Grzyb et al., 2008; Negi et al., 2008). A study performed by Benz et al. (2009) showed that Tic62 is shown to be tightly associated with chloroplast FNR isoforms. However, in vitro experiments performed by Jurić et al. (2009) observed that the interaction between a nuclear encoded TROL
(thylakoid rhodanase-like protein) and FNR is stronger than the interaction between FNR and
35 Tic62.
In the other pathway, ferredoxin is not directly involved (Tagawa et al., 1963; Sazanov et al., 1998a;
Joët et al., 2002; Johnson, 2005). Instead electrons are transferred from NADPH to plastoquinone and studies have shown that a complex, which is similar to the mitochondrial NAD(P)H dehydrogenase (NDH), acts as an electron mediator (Figure 1.6.c). However, several studies pointed out the involvement of Fd in NDH mediated CET pathway (Okegawa et al., 2008; Johnson, 2011,
Yamamoto et al., 2011). NAD(P)H dehydrogenase proteins form a large protein complex within the thylakoid membrane. This complex is known to be involved in both chlororespiration and the CET pathway (Burrows et al., 1998; Sazanov et al., 1998a; Sazanov et al., 1998b). Studies have shown that the concentration of NDH complex involved in this pathway is extremely low and is not compatible with the rate of CET seen in high light conditions (Joliot and Johnson, 2011; Sazanov et al., 1998b). However, chloroplast NDH is essential in the absence of the PGR5 pathway (Munekage et al., 2004; Peng et al., 2011). A study by Takabayashi et al. (2005) showed the involvement of
NDH mediated CET in C4 plants to supply ATP to the CO2-concentrating mechanism.
The presence of NAD(P)H dehydrogenase (NDH) in chloroplasts was first identified by analysing the complete plastid genome sequences in tobacco (Nicotiana tabacum) and liverwort (Marchantia polymorpha) (Shinozaki et al., 1986; Ohyama et al., 1986). 11 genes (ndhA-ndhK) in plastid genome encodes subunits which are homologous to mitochondrial NADH dehydrogenase (complex
1) (Matsubayashi et al., 1987; Peng et al., 2011). Involvement of NDH in CET was first discovered in the M55 mutant in Synechocystis sp. PCC 6803 (Ogawa, 1991). Apart from CET, cyanobacterial
NDH-1 is also involved in CO2 uptake (Mi et al., 1992; Mi et al., 1994; Mi et al., 1995). There are two functionally distinct NDH-1 complexes were found in cyanobacteria. NDH-1L mediates respiration and CET whereas, NDH-1MS mediates CO2 uptake (Ohkawa et al., 2000; Battchikova,
36 2011). NDH-1 in Escherichia coli (E. coli) has 14 subunits. The crystal structure identified in
Thermus thermophilius and E. coli showed the presences of two major parts as membrane and peripheral (Sazanov and Hinchliffe, 2006; Efremov et al., 2010). The membrane segment has 7 hydrophobic subunits with a total of 63 transmembrane helices which are important for proton translocation (Efremov et al., 2010). The peripheral segment has 7 hydrophilic subunits which contain redox centers, flavin mononucleotides and Fe-S clusters (Sazanov and Hinchliffe, 2006).
The structure of the chloroplast NDH is closest to cyanobacterial NDH-1 complex. However, the chloroplast NDH has subunits specific to higher plants (Peng et al., 2011). The chloroplast NDH contains four parts designated as A, B, membrane and lumen. From these subcomplexes, the B and lumen parts have subunits specific to plants (Ishihara et al., 2007; Peng et al., 2008; Peng et al.,
2009; Sirpio et al., 2009a; Suorsa et al., 2010; Yabuta et al., 2010). It was found that, several auxiliary proteins are associated with the chloroplast NDH complex and involved in biogenesis and stability at the transcriptional and post-transcritional level (Rumeau et al., 2007). AtCYP20-2 is one such auxiliary protein associated with the chloroplast NDH complex (Sirpio et al., 2009b). Studies have shown that the chloroplast NDH is associated with multiple copies of PSI to form NDH-PSI supercomplex (Jansson, 1999; Peng et al., 2009). Light harvesting complex 1 (LHCI) proteins,
Lhca5 and Lhca6 are essential for the formation of NDH-PSI supercomplex (Peng et al., 2009; Peng et al., 2011). Recent electron microscopy analysis performed by Kouřil et al. (2014) described two forms of PSI-NDH supercomplexes in barley. The large complex consists of one NDH complex with two copies of the PSI and the small complex has only one PSI associated with one NDH complex. In addition, the pseudo atomic model constructed with the knowledge of the X-ray structure of the bacterial NDH-1 complex (Baradaran et al., 2013) and PSI complex (Amunts et al.,
2010) indicates asymmetric binding of two PSI complex to NDH (Kouřil et al., 2014). This suggested that Lhca5 and Lhca6 subunits mediate the binding of one of the PSI complex to NDH.
37 a Fd b Fd FNR PGR5 1 1 I I I L L
6 C Cyt b f PQ PQ 6 C
R Cyt b f PSI R PSI H H L G G L P P PC
PC H+ H+
Fd c
Cyt b6f PQ NDH PSI
H+ H+
PC
Figure 1.6. Possible pathways of cyclic electron transport around PSI (reproduced from Shikanai,
2014). (a) In Arabidopsis thaliana, PSI forms a complex with PGRL1 and PGR5 which involved in the antimycin A sensitive cyclic electron transport (CET) (DalCorso et al., 2008; Hertle et al.,
2013). (b) The possible pathway of CET in Chlamydomonas reinhardtii with the involvement of the
+ PSI-LHCI-LHCII-Fd-NADP reductase (FNR)-Cyt b6f-PGRL1 supercomplex in State 2. However, the electron flow from Fd to PQ is unclear. This pathway is insensitive to antimycin A (Iwai et al.,
2010) (c) NDH-PSI supercomplex observed in angiosperms (Peng et al., 2009; Ifuku et al., 2011) which transfer electron from Fd to Cyt b6f via plastoquinol. Similar to bacteria and mitochondrial complex I, NDH is involved in generating a proton gradient across the thylakoid membrane
(Baradaran et al., 2013). This figure do not represents the actual positions of each proteins in the complex.
38 1.2.3. The Calvin-Benson-Bassham Cycle
The Calvin-Benson cycle or the light-independent cycle of photosynthesis takes place in the stroma of the chloroplast (Figure 1.7). Products from the electron transport chain, ATP and NADPH are used to drive fixation of CO2 to form sugars. The Calvin-Benson cycle can be divided into three phases: carboxylation of the CO2 acceptor molecule, reduction of 3-phosphoglycerate and regeneration of the CO2 acceptor ribulose 1,5-bisphosphate (RuBP). Carboxylation of RuBP is the first reaction of the Calvin cycle. This reaction results in the formation of two molecules of 3- phosphoglycerate (3-PGA), a reaction catalysed by Rubisco (ribulose-1,5-bisphosphate carboxylase/oxygenase). Reduction of 3-PGA to triose phosphate/glyceraldehyde 3-phosphate includes two steps. The first step is phosphorylation using ATP, catalysed by phosphoglycerate kinase. The second step is the reduction of the product by NADPH producing triose phosphate. This step is catalysed by glyceraldehyde 3-phosphate dehydrogenase. Finally, triose phosphate is either sent out to the cytosol to produce sucrose, incorporated into starch in the chloroplast or used to regenerate RuBP to complete the cycle. For every three molecules of CO2 fixed by Rubisco, six molecules of triose phosphates are generated, one of which can be removed from the cycle.
Equation 1 shows the formation of six molecules of triose phosphate in the carboxylation and the reduction phases of the Calvin-Benson cycle. These reactions use energy (ATP) and reducing equivalents (NADPH) generated in the thylakoid membranes of chloroplasts. Equation 2 shows from six triose phosphates produced, five of them are used in the regeneration phase that restores
RuBP. This reaction uses energy (ATP).
Equation 1:
3CO2 + 3RuBP + 3H2O + 6NADPH + 6ATP 6 triose phosphates + 6NADP+ + 6ADP +6Pi
39 Equation 2:
5 triose phosphates + 3ATP RuBP + 3ADP + 3Pi
Rubisco, catalyzes both the carboxylation and oxygenation of RuBP. RuBP produces two molecules of 3-phosphoglycerate through carboxylation while oxygenation produces one molecule each of 3- phosphoglycerate and 2-phosphoglycolate (Equation 3) (Bowes and Ogren, 1972; Laing et al.,
1974). The oxygenation of RuBP catalyzed by Rubisco initiates a coordinated network of enzymatic reactions in three cellular organelles, chloroplasts, leaf peroxisomes, and mitochondria. This process is known as photorespiration which causes the partial loss of CO2 fixed by the Calvin-Benson cycle
(Jensen, 2000). Increases in photorespiration (oxygenation) relative to photosynthesis
(carboxylation) significantly limits the efficiency of photosynthetic carbon assimilation under unfavourable environmental conditions (Lawlor, 2002; Carmo-Silva et al., 2010).
Equation 3:
+ RuBP + O2 2-phosphoglycolate + 3-phosphoglycerate + 2H
A study by Lorimer et al. (1976) showed that, to activate Rubisco, the active site of the enzyme, the lysine residue in the catalytic site must be carbamylated by an activator CO2 (this is separate from
2+ the substrate CO2) and Mg must bind to the active site before binding RuBP. The enzyme,
Rubisco activase plays a major role, which is involved in the uncoupling of Rubisco and RuBP so
2+ that the site can bind the activator CO2 and Mg in activating Rubisco (Salvucci et al., 1985;
Jensen, 2000; Carmo-Silva and Salvucci, 2013). This catalytic reaction is driven by ATP (Portis,
1995).
40 Light controls the activity of enzymes associated with the Calvin-Benson cycle through the ferredoxin-thioredoxin system (Fd/Trx system) (Bassham et al., 1950; Buchanan, 1980; Buchanan,
1991; Schürmann and Jacquot, 2000; Lemaire et al., 2007; Schürmann and Buchanan, 2008;
Buchanan et al., 2012). The ferredoxin-thioredoxin system consists of ferredoxin (Fd), ferredoxin- thioredoxin reductase, and thioredoxin (Trx) (Schürmann and Buchanan, 2008; Michelet et al.,
2013). Electrons transported through PSI reduces Fd which then activates the enzyme, ferredoxin- thioredoxin reductase. This enzyme then converts a light-activated redox electron signal into a thiol signal which is transmitted to thioredoxin (Schürmann and Buchanan, 2008). Reduced Trx interacts with specific disulfide sites on enzymes of the Calvin-Benson cycle (Foyer and Noctor, 2005;
Schürmann and Buchanan, 2008). In addition to that, the Fd/Trx system is involved in activation of enzymes in different metabolic pathways, such as enzymes which participate in indirect regulation of the Calvin-Benson cycle, in light-dependent ATP production or in diverse carbon metabolism pathways (Lemaire et al., 2007; Schürmann and Buchanan, 2008). This light-dependent regulatory system is only associated with oxygenic photosynthetic organisms (Schürmann and Buchanan,
2008). Studies performed on redox regulation of the Calvin-Benson cycle suggested that all enzymes of the cycle and several associated regulatory proteins may undergo redox regulation through multiple redox post-translational modifications including glutathionylation and nitrosylation (Michelet et al., 2013).
41 3CO2 + 3H2O
6ATP RuBP 6ADP+ 6Pi Rubisco
Ribulose-5-phosphate 3-Phosphoglycerate
3ATP 6NADPH
3ADP+ 3Pi 6NADP +6Pi
Triose phosphate 1,3-bisphosphoglycerate
Sucrose/starch production
Figure 1.7. A diagram representing the major steps in the Calvin cycle or the light-independent
reactions. three molecules of CO2 and three molecules of water are combined with RuBP to produce two 3-carbon molecules. 3-phosphoglycerate (3-PGA) catalysed by Rubisco. ATP and NADPH produced in light reaction are consumed during the phosphorylation and the reduction of 3-PGA.
Triose phosphate is either sent out to cytosol to produce sucrose, incorporated into starch in the chloroplast or used to regenerate RuBP to complete the cycle. Oxygenation of RuBP is not indicated in this figure.
42 1.3. Effects of abiotic or environmental stress on plants
Plants are frequently exposed to combinations of stresses. Abiotic stresses, including low temperature, salinity, drought, flooding, heat, oxidative stress and exposure to heavy metals, and biotic stresses including infections from pathogens like bacteria, virus, fungi, herbivores are responsible for severe crop losses every year (Mahajan and Tuteja, 2005). Stress can be defined as an adverse force or a condition, which inhibits the normal functioning and well being of a biological system such as plants (Levitt, 1980; Jones and Jones, 1989). Abiotic stresses can be responsible for the inhibition of photosynthesis and decreases in crop productivity (De Oliveira et al., 2013).
1.3.1. Salt Stress
Salinisation is the accumulation of soluble salts including sodium, magnesium and calcium in the soil, which decrease the fertility of that soil (European Soil Portal, 2012). Salt affected lands can be found in all climatic regions, from the humid tropics to the polar regions. Saline soil is also found in places below sea level, such as the area around the Dead Sea and in high mountainous regions, such as the Tibetan Plateau and the Rocky Mountains (Pitman and Läuchli, 2002; Manchanda and Garg,
2008). Salinity can be classified as primary or secondary. Primary salinity is the accumulation of soluble salts over a long period of time through weathering of rocks containing salts and deposition of salts from oceans carried by wind and rain (Manchanda and Garg, 2008). Secondary salinity is the accumulation of soluble salt in the surface of the soil, due to rises in the water table by the exploitation of the land and water through agricultural practices (Manchanda and Garg, 2008). Soil salinity can also be categorized as either irrigated land salinity or dry land salinity. Irrigated land salinity is the rise of salt-contaminated ground water and accumulation of soluble salt in crop fields.
43 The major causes of irrigated land salinity are over-irrigation of farm lands, inefficient water use, poor drainage and irrigation on unsuitable soils which make salt 'leak' and become deposited in water channels, drains and water stores (Irrigation salinity, 2011). Dry land salinity mainly occurs in the arid regions of the world, where salt moves to the soil surface and concentrates due to evapotranspiration, making the land unsuitable for agricultural purposes (Salinity Factsheet, 2011).
Salinity is one of the major abiotic stresses which cause crop losses every year and it is predicted to get considerably worse over the next 30-50 years (Boyer, 1982; Nelson et al., 1998; Tuteja, 2007).
Salt in soil affects photosynthetic organisms including plants, algae and cyanobacteria in two ways.
First, salt triggers osmotic stress in plants or other photosynthetic organisms, due to low water availability resulting from the negative water potential of the soil. Second, ionic stress occurs through solute imbalance, due to the high levels of Na+ and Cl- in the cytosol and changes to the intracellular K+/Na+ ratio (Blumwald et al., 2000; Conde et al., 2011). Research has been carried in to identify genes responsible for stress tolerance in photosynthetic organisms. Stress tolerance is a multigenic trait, therefore it is difficult to understand the involvement of genes in stress tolerance through a gene-by-gene approach (Brinker et al., 2010). Projects, including complete genome sequencing and large-scale expressed sequence tag (EST) sequencing have been performed to identify gene functions in stress tolerance (Ahmad et al., 2013). Work from Hatzimanikatis et al.
(1999) discussed how the interaction between protein abundance and mRNA levels is important to understand gene expression and protein function in stress tolerance. Many genes and proteins including ferritin, HSPs (heat stress proteins), FtSH (the ATP-dependent integral membrane protease), GST (glutathione S-transferase) and proteasome proteins responsible for stress responses and tolerance have been identified through proteomic and transcriptomic analysis (Ahmad et al.,
2013).
44 1.3.1.1. Effects of salt stress on plants
High levels of salt ions, such as Na+, Mg2+and Cl- in soil and water affect plants by disturbing the water uptake from roots, ion homeostasis and causing toxicity (Parida and Das, 2005). Salt stress affects all the major metabolic processes in plants, including protein production, growth, photosynthesis and lipid metabolism (Parida and Das, 2005). Stress responses of plants can divided into two phases as osmotic and toxic (Munns and Tester, 2008). During the osmotic phase, plants show rapid responses to the osmotic effects of salt in soil. The toxic phase is where plants show slower responses due to the accumulation of high levels of Na+ in leaves (Munns and Tester, 2008).
This section aims to introduce some of the short-term and long-term effects of salt stress on plants.
1) Effects of salt on plant growth
Salinity restricts water uptake by roots, stunting plants due to reduced cell expansion (Zhu, 2002;
Parida and Das, 2005). Excess salt reduces the fresh and dry weight of leaves, stem and roots
(Hernández et al., 1995; Takemura et al., 2002). Increased levels of salt ions around roots causes loss of cell volume and turgor, which leads to a drop in cell elongation of leaves and stems (Yeo et al., 1991; Passioura and Munns, 2000; Cramer, 2002; Fricke and Peters, 2002). Decreases in the rate of surface expansion, which can lead to the complete interruption of leaf expansion, is an immediate response to the high concentrations of salt ions in plant cells (Wang and Nii, 2000). A study has shown that an 80% of growth reduction occurred in Raphanus sativus (radish) is due to the decrease in the leaf area expansion, which drops light interception and 20% is due to the stomatal closure
(Marcelis and Hooijdonk, 1999). A study by Mohammad et al. (1998) showed that in tomatoes, shoot weight, plant height, number of leaves per plant, root length and root surface per plant decreased with increasing salt concentrations. Similar results were observed in cotton (Meloni et al.,
45 2001) and Brassica campestris sp. chinensis (Memon et al., 2010) when exposed to high salt concentrations. However, the exact mechanism of the downregulation of leaf and shoot growth under stress is not known (Munns and Tester, 2008). Limitations in photosynthesis may cause long term effects on the growth rate in plants under salinity (Zhu, 2001; Munns and Tester, 2008).
Phytohormones such as abscisic acid (ABA) and gibberellic acid (GA) play a major role in the regulation of shoot and root growth under salt stress (Spollen et al., 2000; Sharp and LeNoble,
2002; Achard et al., 2006; Munns and Tester, 2008). A study by Achard et al. (2006) showed that salt induces ABA and ethylene signalling pathways and regulates the growth of plants through activating functions of DELLA proteins (DELLA proteins are nuclear proteins which restrict the cell proliferation and expansion). A recent study by Duan et al. (2013) showed that, lateral root growth in Arabidopsis thaliana (A. thaliana) is regulated through the ABA signalling pathway which originated in the endodermis. Further, they showed that the endodermis is the point of cross talk between ABA and GA pathways (GA antagonises the ABA pathway), which cause the regulation of root growth under salt stress.
2) Effects of salt on photosynthesis
Photosynthesis is one of the major physiological processes affected by salt stress. Salinity in soil either causes short-term or long-term effects on photosynthesis (Parida and Das, 2005). Salinity in soil prevents water uptake by plants, which results in drought stress. This causes stomatal closure in leaves to reduce water release from transpiration (Hsiao, 1973; Fricke et al., 2004; Munns and
Tester, 2008; Negi et al., 2014). Stomatal closure under salinity occurs due to loss of leaf turgor which is generated through either high vapour pressure deficit or a ABA mediated signal (Munns and Tester, 2008; Chaves et al., 2009). As a result, photosynthesis is inhibited, due to low CO2 assimilation. Although low photosynthetic rates initially occur due to the stomatal closure, as salt
46 stress becomes more severe, photosynthesis is inhibited due to metabolic impairments (Cornic et al.,
1989; Sharkey, 1990; Cornic and Briantais, 1991; Panković et al., 1999). In addition to reduced CO2 diffusion through stomata, salt restricts CO2 diffusion through the leaf mesophyll (gm) (Flexas et al.,
2004; Flexas et al., 2007). This might be due to several reasons, like physical alterations in the structure of intercellular spaces caused by leaf shrinkage (Lawlor and Cornic, 2002) or changes in the biochemistry or membrane permeability (Gillon and Yakir, 2000; Flexas et al., 2008). A study by Kao et al., (2001) pointed out that photosynthetic rate and stomatal conductance were decreased in seedlings of a mangrove species, Kandelia candel when salt concentration increased up to 430 mM. Work from Stepien and Johnson (2009) showed that salt affects PSII and PSI photochemistry and the total leaf chlorophyll content in A. thaliana when exposed to high salt concentrations while the halophyte, Thellungiella salsuginea (previously Thellungiella halophila) was unaffected.
3) Effects of salt on water relations and ion balance in plants
Although the turgor pressure of plant cells increases, both osmotic and water potential drop with increasing salinity (Morales et al., 1998; Hernández et al., 1999; Khan et al., 1999; Meloni et al.,
2001; Aziz and Khan, 2001; Romero-Aranda et al., 2001). Matsumura et al. (1998) showed that in both Chrysanthemum and sea aster, the leaf osmotic potential decreased with increasing salt.
Similar to that, another study on a halophytic perennial grass, Urochondra setulosa showed a drop in water potential, water uptake, transpiration rate, water retention and water use-efficiency under salt stress (Gulzar et al., 2003). Unlike glycophytes, halophytes do not show significant change in the osmotic or water potential under salt stress. A study on Rhizophora mucronata showed that in leaves, the water potential, osmotic potential and xylem tension increased with increasing salt (Aziz and Khan, 2001). Although leaf relative water content is unchanged in Suaeda salsa, the water potential and evaporation rate decreases with increasing salt concentration (Lu et al., 2002). Salt
47 (NaCl) affects the ion uptake of plants. Studies have shown that, under high salt concentrations, Na+ and Cl- accumulated in plants decrease the uptake of other ions including Ca2+, K+ and Mg2+ (Khan et al., 1999; Khan et al., 2000; Rengasamy, 2010). Apart from that, excess Na+ in plants inhibits the activity of many enzymes in plants (Zhu, 2002). A study has shown that compared to roots and stem, leaves accumulate more Na+ and Cl- than K+ and Mg2+ (Ferreira et al., 2001). Another study on barley showed that high concentrations of Na+ reduce the uptake of Ca2+ and K+ and decrease photosynthesis by reducing the stomatal conductance whereas, Cl- in leaves affects photosynthesis through non-stomatal effects including, chlorophyll degradation and decreasing the efficiency of excitation energy capture (Tavakkoli et al., 2010).
4) Effects of salt on photosynthetic pigments, proteins and lipid composition
Salt has a major impact on photosynthetic pigments, such as chlorophylls and carotenoids in leaves and this causes leaf chlorosis and senescence (Hernández et al., 1995; Hernández et al., 1999;
Gadallah, 1999; Agastian et al., 2000). Work from Kennedy and DeFillippis (1999) pointed out that, compared to chlorophyll-a and carotenoids, the decline of protochlorophylls and chlorophyll-b is greater with increasing salt. Another study on pigment composition in tomato showed that salt affects the total chlorophyll and carotene content in leaves (Khavari-Nejad and Mostofi, 1998).
Studies have shown a change in the protein and lipid composition of plants under salt stress. Work from Parida et al. (2002) showed that leaf protein content decreased with increasing salt in
Bruguiera parviflora. A study by Hassanein (1999) pointed out that peanut (Arachis hypogaea L.) exposed to high NaCl concentration showed an increase in certain polypeptides (127 and 57 kDa) and a decrease in others (260 and 38 kDa). Lipids are important as they act as an efficient energy store, hormones and as a part of the cellular membranes (Singh et al., 2002). The phospholipid
48 bilayers are involved in mechanisms of desiccation tolerance in plants. Sugars like trehalose help to stabilize the lipid bilayers under water deficit conditions (Singh et al., 2002). Other than that, increases in unsaturated fatty acids in plant cell membranes acts against drought and salt stress
(Singh et al., 2002; Sui et al., 2010). Salt decreased the molar percentage of sterols and phospholipids in the root plasma membrane in the salt marsh grass species, Spartina patens (Wu et al., 1998). However, the ratio of sterol and phospholipids is unaffected. Excess salt in soils increases cell respiration and has been seen to cause membrane instability and changes in membrane permeability (Gupta et al., 2002).
5) Effects of salt on leaf anatomy and the structure of chloroplast
Increases in the epidermal thickness, mesophyll thickness, palisade cell length and diameter and spongy cell diameter are major long term effects of salt on leaf cell anatomy (Longstreth and Nobel,
1979). A study on the leaves of sweet potatoes showed that salinity also initiates vacuolation development, swelling in the endoplasmic reticulum and mitochondria, decline in mitochondrial cristae, forms vesicles and fragments in the tonoplast and causes degradation of the cytoplasm
(Mitsuya et al., 2000). Apart from that, studies have shown that salt affects the size of intercellular spaces in leaves, decreases the number of chloroplasts in leaves and causes rounding of cells (Bruns and Hechtbuchholz, 1990; Delfine et al., 1998; Parida et al., 2004). Salt also affects the stomatal density of leaves (Romero-Aranda et al., 2001). Electron microscopy studies on the thylakoid structure showed that the thylakoids became disorganized and the starch content in cells decreased when exposed to salt (Hernández et al., 1995; Hernández et al., 1999). Bruns and Hechtbuchholz
(1990) showed that, the amount and the depth of grana stacks decrease with increasing salt. Apart from that, their study also pointed out that salt affects the ultrastructure of the chloroplasts due to the enlargement of the thylakoids and starch grains (Bruns and Hechtbuchholz,1990).
49 6) Effects of salt on the nitrate and malate metabolism
Nitrate reductase (NR) is one of the most important enzymes of the nitrate assimilation pathway of
- - plants. High levels of Cl in leaves interrupt NO3 uptake which leads to the drop in NR activity of
Zea mays (Flores et al., 2000; Abd-El Baki et al., 2000). Similar results have been seen in studies performed on different plants, including beans (Gouia et al., 1994), sugar beet (Ghoulam et al.,
2002) and tomatoes (Debouba et al., 2007). However, Reda et al. (2011) showed that NR activity actually increased when cucumber roots were exposed to 200 mM of NaCl for 60 minutes. This suggests that the salt induced modification of NR activity depends on nitrogen source, species, salt concentration and the exposure time (Reda et al., 2011). Salt also disrupts the nitrogen fixation of plants by reducing nodulation and nitrogenase activity (Soussi et al., 1999). A study by Serraj et al.
(1998) pointed out, that in legumes such as soybean, common bean and alfalfa a rapid decrease in growth occurred when nodulated roots were exposed to NaCl. In higher plants, NADP-malate dehydrogenase (NADP-MDH) in chloroplasts converts oxaloacetate to malate and undergoes rapid reversible light activation (Ferte, 1986; Buchanan, 1991; Jacquot et al., 1997; Scheibe, 2004).
Several studies have shown changes in the levels of NADP-MDH under salt stress. A study by
Cushman (1993) showed a significant change in NADP-MDH transcripts in leaves compared to roots under salinity, suggesting the involvement of NADP-MDH in the CO2 fixation pathway of
CAM (Crassulacean acid metabolism) in the facultative CAM plant, Mesembryanthemum crystallinum. Another study has shown that, after three weeks of salt treatment high levels of Na+ were found in the shoots of Eucalyptus citriodora (Aragao et al., 1997). However at this stage, growth of plants is not affected by salt and a decrease in malate content and an increase in the activity of NAD and NADP-malic enzymes were observed.
50 1.3.1.2. Sensing and signal transduction in salt stress tolerance
Elevated levels of Na+ in the soil solution cause hyper osmotic stress in plant roots. Na+ influx in roots occurs through voltage-dependent non-selective cation channels (NSCC) and possibly other cation channels (Figure 1.8) (Tester and Davenport, 2003; Horie and Schroeder, 2004). Changes in osmotic pressure in roots are detected through plant hyperosmotic sensors and Na+ sensors in roots
(Bartels and Sunkar, 2007; Deinlein et al., 2014). However, these sensors have remained elusive in plants. SLN1 and SHO1 are the two osmosensors, important for the HOG (high osmolarity glycerol)-MAPK (mitogen-activiated protein kinase) pathway in yeast (Posas et al., 1998; Reiser et al., 2003). A study by Urao et al. (1999) showed that a transmembrane hybrid type histidine kinase
(HK1) from A. thaliana acts as an osmosensor in SLN1-defective yeast mutants.
In contrast to signal perception, several signal transduction pathways have been identified in plants
(Bartels and Sunkar, 2007). These involve of networks of protein-protein interactions and signalling molecules such as Ca2+ and reactive oxygen species (ROS) (Bartels and Sunkar, 2007). MAPKinase and SNF1/AMP activated protein kinase pathways are examples for such signal transduction processes involved in stress regulation in plant (Morrison and Davis, 2003; Bartels and Sunkar,
2007). The MAPK pathway is involved in signal transduction through protein phosphorylation
(Posas et al., 1998; Morrison and Davis, 2003). Three protein kinases: MAPKKK (MAPK Kinase
Kinase) activates MAPKK (MAPK Kinase) by phosphorylation of specific serine/threonine residues and MAPKK activates MAPK by phosphorylation of tyrosine and threonine residues
(Posas et al., 1998). MAPK in the cytoplasm translocate into nucleus where kinase activates genes through phosphorylation of transcription factors (Treismann, 1996). MAPK is also involved in phosphorylation of specific enzymes and cytoskeleton components of cells (Robinson and Cobb,
1997). SNF1/AMP-activated protein kinase activates through the phosphorylation of
51 serine/threonine residues (Halford and Hardie, 1998). These kinases sense ATP/AMP ratios in cells and regulate genes involved in carbohydrate metabolism (Hardie et al., 1998; Halford and Hardie,
1998). Some SNF1 kinases are expressed in response to ABA or dehydration of cells (Kobayashi et al., 2004). Apart from these protein kinases, phosphotases, phospholipids, salicylic acid and nitric oxide are involved in the osmotic stress signalling (Bartels and Sunkar, 2007). Hyperosmotic sensors or Na+ sensors in root cells interact with Ca2+channels to increase the levels Ca2+ in the cytosol under salinity (Knight et al., 1997; Tracy et al., 2008; Kiegle et al., 2000; Martí et al., 2013;
Kurusu et al., 2013). Ca2+ act as a second messenger in the stress regulation pathways in plants
(Snedden and Fromm, 1998; Snedden and Fromm, 2001). Three major class of Ca2+ sensors were found in plants. They are calmodulin, Ca-dependent protein kinases (CDPKs) and calcineurin B- like proteins (CBLs) with CBL-interacting protein kinases (CIPKs) (Yang and Poovaiah, 2003;
Weinl and Kudla, 2008). All of these Ca2+ sensors are involved in stress signal transduction in plants
(Snedden and Fromm, 2001; Luan et al., 2002; Zhu, 2000). These Ca2+-dependent kinases may transduce hyperosmotic signals which regulates protein activity and the gene transcription in cells
(Deinlein et al., 2014).
Transcription of stress-induced genes is a complex process driven by two components: transcription factors and their associated cis-regulatory elements (CREs) (Gómez-Porras et al., 2007).
Transcription factors are proteins which play a major role in regulation of gene expression under salinity and act as a link which combines salt sensory pathways to stress tolerance mechanisms in plants (Deinlein et al., 2014). Plants contain large number of transcription factors involved in transcriptional regulation which is necessary for plant development and stress regulation (Bartels and Sunkar, 2007). Riechmann et al. (2000) showed that, in A. thaliana, over 5% of the genome is devoted to encoding more than 1500 transcription factors. These can be classified into families such as basic leucine zipper (bZIP), WRKY, APETALA2/ETHYLENE RESPONSE FACTOR
52 (AP2/ERF), MYB, basic helix-loop-helix (bHLH) and NAC (Kasuga et al., 1999; Tran et al., 2004;
Yang et al., 2009; Jiang and Deyholos, 2009; Jiang et al., 2009; Cui et al., 2013). CREs are mainly located in the core promoter of a gene and categorized as the ABA-responsive element (ABRE) and the dehydration responsive element (DRE) (Baker et al., 1994). ABA is known to be involved in gene regulation mainly at the transcription level (Busk and Page, 1998; Chak et al., 2000). These
ABRE or DRE consists of certain nucleotide sequences which can be identified by transcription factors (Gómez-Porras et al., 2007). For example, the ACGT core of a motif in the promoter of a
ABA-responsive gene is recognized by plant bZIP proteins (Guiltinan et al., 1990; Hattori et al.,
1995; Hobo et al., 1999; Choi et al., 2000; Uno et al., 2000).
After stress induction, one of first responses of the salt tolerance mechanisms in plants is the regulation of growth which is correlated with the changes of the levels of phytohormones including,
ABA, jasmonates, gibberellic acid and brassinosteroids (Kilian et al., 2007; Geng et al., 2013).
Plant stress tolerance mechanism consists of three components: (1) osmotic tolerance, which maintains water uptake by roots and growth regulation (2) Na+ exclusion with the involvement of various Na+ transporters and ion channels (3) tissue tolerance through compartmentalization of excess Na+ into vacuoles and other tissues, accumulation of compatible solutes, activation antioxidant system and plant hormones (Munns and Tester, 2008).
53 Cellular influx Sensing and signalling Transcriptional Control
? Apoplast
NSCC others ?
cytosol CBLS Nucleus
CIPKs WRKY bHLH Na+ sensors? Ca+ Omotic sensors Xylem Loading Na+ ? NAC MYB ROS CDPKs bZIP KORC AP2/ /NORC? ERF Na+
Hormones (ABA) HKT Na+ Na+ exclusion H+ from leaves NHX Osmoprotective Vacuole Osmolytes Na+ proteins
H+
SOS2 Xylem P SOS3 SOS1
Na+
Detoxification Mechanisms
Figure 1.8. Cellular Na+ transport mechanisms and important components of the salt stress responses in plant root cells (reproduced from Deinlein et al., 2014). Na+ entry occurred through NSCC (non-selective cation channels) or though other unknown cation channels. Na+ is sensed by yet unidentified sensors in the cytosol. Ca2+, ROS and hormone signalling pathways (Calmodulin, Ca-dependent protein kinases
(CDPKs) and calcineurin B-like proteins (CBLs) with CBL-interacting protein kinases (CIPKs) are part of
Ca2+ signalling pathway) activate transcription factors in the nucleus which are important for the salt- induced gene expression. These activates cellular detoxification through activating Na+ transporters including, NHX, SOS1 pathway and HKT1 and cause accumulation of osmolytes and osmoprotective proteins. KORC and NORC are the outward-rectifying ion channels. 54 1.3.1.3. Transcriptomics and proteomics of salt tolerance
Salt tolerance in plants is complex and controlled by many genes and biochemical-physiological mechanisms (Ciarmiello et al., 2011). Salt-responsive genes can be classified into several functional groups (Ciarmiello et al., 2011) (Table 1.1). Many studies have been performed on Arabidopsis and crops to understand the mechanisms and the signalling pathways of salt tolerance (Zhang and Shi,
2013). Genomics, transcriptomics, proteomics, metabolomics and ionomics have been extensively used to identify abiotic stress tolerance in plants (Cramer et al., 2011).
Microarrays are one of the powerful techniques used for genomic wide transcript expression profiling in many plants, including Arabidopsis thaliana, Vitis vinifera and Hordeum vulgare under stress (Kreps et al., 2002; Walia et al., 2006; Cramer et al., 2007; Shelden et al., 2013). Microarrays have also been used to analyse cell-type specific transcripts in plants such as A. thaliana, maize, rice, barley and soybean (Pu and Brady, 2010; Long, 2011; Rogers et al., 2012). RNA-Seq or whole transcriptome shotgun sequencing is another useful technology which uses the capabilities of next- generation sequencing (NGS) and provides a strategy to identify and quantify changes in the transcriptomes (Shelden et al., 2013). Compared to microarrays, NGS technology is more useful as it used for the gene expression profiling in many plant species including models such as A. thaliana
(Weber et al., 2007) and Thellungiella parvula (Dassanayake et al., 2011) and agronomic species like soybean (Fan et al., 2013), Lolium perenne L. (Studer et al., 2012), Zea mays (Li et al., 2010),
Sorghum bicolor (Dugas et al., 2011), Panicum virgatum L. (Wang et al., 2012) and Triticum aestivum (Gillies et al., 2012). Apart from that, NGS provides better quantitation and accuracy than microarrays (Jain, 2011). NGS analysis has been performed on plants to examine abiotic stress responses, including salinity (Molina et al., 2008; Fan et al., 2013), cold stress (Tamura and
Yonemaru, 2010) and drought (Dugas et al., 2011; Dong et al., 2012; Vidal et al., 2012). Using
55 RNA-Seq and the sorghum genome sequence, Dugas et al. (2011) identified over 28,000 genes which are transcriptionally regulated in response to osmotic stress and ABA.
Table 1.1. Functional groups of genes/proteins induce under salt stress
Functional groups of Products Example References genes/proteins activated under salinity 1) Genes encoding Heat-stress Physcomitrella patens Ruibal et al., 2013 products which directly proteins PpHsp16.4 protect plant cells (HSP)/Chaperones under stress Late Barley HVA1 overexpressed Xu et al., 1996 Embryogenesis in rice Abundant (LEA) proteins Osmoprotectants Glycine betaine Chen and Murata, 2011 Detoxification Pisum sativum ascorbate Hernández et al., enzymes and free peroxidase, glutathione 2000 radical scavengers reductase, monodehydroascorbate reductase, dehydroascorbate reductase, superoxide dismutase
2) Genes encoding Mitogen-activated Arabidopsis thaliana Mizoguchi et al., products which induce protein kinase 1996 AtMPK3 signalling pathways and (MAPK) transcriptional control Ca-dependent Dubrovina et al., protein kinase Vitis amurensis CDPK 2013 (CDPK) SOS-kinase Arabidopsis thaliana SOS3- Xiong et al., 2002 SOS2 protein kinase Phospholipases Arabidopsis thaliana non- Peters et al., 2014 specific phospholipase C5 (NPC5) Transcriptional factors Medicago truncatula MtCBF4 Li et al., 2011 3) Genes encoding Aquaporins Tobacco NtAQP1 Sade et al., 2010 products which involve in Ion transporters Wheat Laurie et al., 2002 water and ion uptake and K+/Na+ transporter HKT1 transport
56 'Omic' studies have provided information on gene expression at the mRNA level in many plants
(Zhang et al., 2001; Wang et al., 2004; Wong et al., 2005; Wong et al., 2006; Zouari et al., 2007;
Zhang et al., 2008; Diédhiou et al., 2009; Jha et al., 2009). Studies found that more than 194 transcripts in A. thaliana, 10% of transcripts in salt-tolerant rice and 2300 ESTS/cDNAs in some halophytes such as, Thellungiella halophila, Suaeda salsa, Aeluropus littoralis, Salicornia brachiata and Festuca rubra sp. were expressed in response to salinity (Zhang et al., 2001; Wang et al., 2004; Wong et al., 2005; Wong et al., 2006; Zouari et al., 2007; Zhang et al., 2008; Diedhiou et al., 2009; Jha et al., 2009). However, due to post-transcriptional events and post-translational modifications (phosphorylation and glycosylation) mRNA levels do not necessarily correlate with the expression levels of proteins. Therefore, it is important to study salt stress responses of plants at protein levels. Combination of protein chromatography, proteolytic digestion and peptide mass spectroscopic analysis were used to characterize proteomes effectively (Aebersold and Goodlett,
2001; Mann et al., 2001). Many proteomic studies were performed to identify salt-responsive proteins in plants (Lee et al., 2004; Ndimba et al., 2005; Jiang et al., 2007; Zhang et al., 2009;
Tanou et al., 2009; Rasoulnia et al., 2011). According to a review published by Zhang et al. (2012)
2171 salt-responsive proteins have been identified from leaves, roots, shoots, seedlings, unicells, grains, hypocotyles, radicles and panicles from 34 plant species. Another study showed the differences of salt responsive proteomes of salt-sensitive and salt-tolerance plants expressed under salinity (Kosová et al., 2013). Table 1.2 contains some examples for genes and proteins expressed under salt stress.
Table 1.2. Selective examples of genes/proteins induced by salt stress
Plant species Genes/proteins Characteristic features References Oryza sativa MYB (OsMYB2) Accumulation of soluble Yang et al., 2012 sugars and proline SKC1 Decrease Na+ concentration Ren et al., 2005
57 in xylem and shoots HKT (OsHKT2) Na+ transporter Horie et al., 2007; 2011 OtsA, OtsB Better plant growth and Garg et al., 2002 decrease photooxidative damage TPS, TPP Increase growth Jang et al., 2003 performance and photosynthetic capacity Arabidopsis Sal1 Helps to overcome Na+ and Quintero et al., 1996 thaliana Li+ toxicity NHX1 Na+ compartmentalization Apse et al., 1999 SOS1 Better root growth, PSII Shi et al., 2003 activity and survival under salt stress conditions
CodA Better germination and Hayashi et al., 1997 photosynthetic activity ALDH3 Reduce lipid peroxidation Sunkar et al., 2003 P5CS Catalyze the rate limiting Liu and Zhu, 1997 step of proline synthesis GSK1 Increase root growth, Piao et al., 2001 expression of salt- responsive genes, ATCP1, RD29A, ATCBL1 Triticum aestivum Nax1, Nax2 Control Na+ accumulation James et al., 2006 and tolerance Brassica napus Bnd22 22KDa protein, increased by Reviron et al., 1992 salt and water stress Hordeum vulgare hva1 Induced due to stress like Hong et al, 1992 drought, salt and heat Lycopersicon TAS-12 Lipid transfer proteins Torres-Schumann et al., esculentum induced by salt and water 1992 stress Nicotiana tabacum Osmotin 26-KDa protein, induced by Singh et al., 1987 salt and PEG-induced drought Invertase Better photosynthetic Fukushima et al., 2001 capacity Vitis vinifera OEE2 Involved in oxygen- Vincent et al., 2007 evolving in photosynthesis Glycine max Calreticulin Involve in Ca signalling Sobhanian et al., 2010 pathway Zea mays 14-3-3 proteins Involved in salt induced Zörb et al., 2010
58 responses Thellungiella sp CBL9 Encodes calcineurin-B-like Pang et al., 2010 protein which involve in Ca signalling pathway Solanum Peroxidase ROS scavenging enzymes Manaa et al., 2011 lycopersicum Mesembryanthemu H+-ATPases Involved in ion Barkla et al., 1995 m crystallinum homoeostasis Sorghum bicolor Lectins Carbohydrates binding Swami et al., 2011 proteins involved in defence system Salicornia 1,3,4-triphosphate 5/6- Involved in the Wang et al., 2009 europaea kinases phosphorylation of the transcription factors Arachis hypogaea Pathogenesis-related Involved in signalling Jain et al., 2006 proteins pathways under salt stress Aeluropus Voltage-dependent anion Involved in osmotic Sobhanian et al., 2010 lagopoides channel proteins regulation under salt stress Dunaliella salina Nitrate (gi52789941) and Increase the adaptation Katz et al., 2007 ammonium (BM446979) mechanisms under salt transporters
1.3.1.4. Salt tolerance mechanisms in plants
Plants develop various physiological, biochemical and molecular mechanisms to survive in the changing environmental conditions. This section provide a detailed description on known salt tolerant mechanisms of plants.
1) Ion Regulation and Compartmentalization
Ion regulation and compartmentalization is one such method use by both glycophytes and halophytes (Parida and Das, 2005). Accumulation of excess sodium ions in plant cells cause detrimental effects on plant metabolism (Zhu, 2002; Tester and Davenport, 2003; Horie and
59 Schroeder, 2004; Apse and Blumwald, 2007). Regulation of the expression and the activity of K+ and Na+ transporters and H+ pumps are important for the ion compartmentalization as they help to maintain high levels of K+ and low levels of Na+ in the cytosol under salt stress (Zhu et al., 1998;
Schroeder et al., 2013). Na+ transporters are involved in the extrusion of Na+ from cells at the plasma membrane via Na+/H+ antiports, sequestration of Na+ into plant vacuoles and blockage of
Na+ over-accumulation in leaves (Blumwald and Poole, 1985; Shi et al., 2000; Mäser et al., 2002). A tonoplast-localized Na+/H+ exchanger (NHX1) and a plasma membrane-localized Salt Overly
Sensitive 1 (SOS1/NHX7) are two major Na+ transporters, important for the salt stress resistance mechanisms in plants (Blumwald et al., 2000; Qiu et al., 2002; Brini and Masmoudi, 2012;
Yamaguchi et al., 2013). In addition, plasma membrane-localized Na+ transporters, HKT (high affinity potassium transporters) involved in Na+-selective transport or K+- Na+ co-transport in plant cells (Rubio et al., 1995; Uozumi et al., 2000; Mäser et al., 2002). NHX1 is involved in the Na+ detoxification by removing excess Na+ from the cytosol and compartmentalization in the vacuoles whereas SOS1 is important to export Na+ out of the cells (Apse et al., 1999; Deinlein et al., 2014).
HKT1 proteins are important as they involved in the Na+ ascending from xylem sap and recirculating sodium ions from leaves to roots (Ren et al., 2005; Brini and Masmoudi, 2012).
Na+/H+ antiporter, SOS1 is regulated by the SOS2 Ser/Thr protein kinase and two calcium sensors,
SOS3/CBL4 (Calcineurin B-Like 4) and SCaBP8/CBL10 (SOS3 homolog SOS3-Like Calcium
Binding Protein8/ Calcineurin B-Like 10) (Qiu et al., 2002; Quintero et al., 2002; Quan et al.,
2007). This SOS2-SOS3 complex phosphorylates and activates the SOS1 transporter, which extrudes excess Na+ from the cytosol (Qiu et al., 2002; Quintero et al., 2002; Quintero et al., 2011).
60 Activity of SOS1 in ion homeostasis was demonstrated first in A. thaliana (Shi et al., 2000; Shi et al., 2002). Shi et al. (2002) showed that SOS1 is expressed in the epidermis of the root tip region and in xylem parenchyma cells. Studies pointed out that activity of SOS1 coordinates with the activity of HKT1 in the plasma membrane of xylem parenchyma cells to achieve the adequate partition of Na+ between roots and shoots (Sunarpi et al., 2005; Pardo et al., 2006; Pardo, 2010).
Apart from that, it has been found that SOS1 plays a major role in salt tolerance in other plants including, Thellungiella salsuginea, tomato and the moss, Physcomitrella patens (Oh et al., 2009;
Olías et al., 2009; Fraile-Escanciano et al., 2010). SOS1 is known to be involved in Na efflux at the root epidermis and indirectly important for K+ uptake in the cells (Shi et al., 2002; Olías et al.,
2009; Huang et al., 2012). It is also responsible for the ion concentration of cells by controlling Na+ loading and unloading from xylem vessels and long distance Na+ transport from roots to shoots (Shi et al., 2002; Olías et al., 2009). A study by Quintero et al. (2011) showed that the SOS2-SOS3 complex up-regulates SOS1 activity through phosphorylation of the auto-inhibitory domain in
SOS1. The current model of the SOS pathway explains that increase in the intracellular Ca2+ due to high Na+ concentrations encourage the Ca2+ binding to SOS3 which interacts with and activates
SOS2. SOS2 and SOS3 physically interacts and forms the SOS2-SOS3 complex where activated
SOS2 phosphorylates the plasma membrane-localized SOS1. Phosphorylated SOS1 increases the
Na+ efflux under salt stress (Zhang and Shi, 2013). A recent study of Feki et al. (2014) showed that over-expression of a truncated form of wheat SOS1 (TdSOS1, deletion of the auto-inhibitory domain) in A. thaliana enhanced the root elongation, water retention and salt tolerance.
A transporter called NHX1 belongs to the CPA1 family (a monovalent cation/proton antiporter family) (Mäser et al., 2001) and is found on the plasma membrane, in endosomal compartments and in vacuoles (Apse et al., 1999; Shi et al., 2000; Yokoi et al., 2002; Apse and Blumwald, 2007;
61 Rodriguez-Rosales et al., 2008; Hamaji et al., 2009; Leidi et al., 2010; Bassil et al., 2011b).
AtNHX1 over-expression studies by Apse et al. (1999) and Zhang and Blumwald (2001) showed that NHX1 played a major role in salt tolerance in transgenic plants including A. thaliana, tomato and rice. Apart from the sequestration of Na+ within the vacuoles, it is also important for compartmentalization of K+ into the vacuoles and for promotion of cellular pH homeostasis
(Barragán et al., 2012). NHK1 participates in the movement of Na+ or K+ out of the cells or lumenal movement of Na+ or K+ into the vacuoles and intracellular organelles (Bassil et al., 2011b). This electroneutral exchange of Na+ and/or K+ for H+ is driven through the electrochemical gradient generated by proton pumps including, (H+)-ATPase in the plasma membrane and (H+)-ATPase and
(H+)-PPase in the plasma membrane (Sze, 1983; Scherer and Martiny-Baron, 1985; Blumwald and
Poole, 1985; Blumwald and Poole, 1987; Blumwald et al., 2000; Rodriguez-Rosales et al., 2009).
Six intracellular NHX-type antiporters were found in A. thaliana and these can be classified into two groups (Brett et al., 2005; Pardo et al., 2006; Rodriguez-Rosales et al., 2009). NHX1 to NHX4, antiporters associated with vacuoles belong to Group 1 whereas, NHX5 and NHX6 which are associated with endosomal components, belong to Group 2 (Bassil et al., 2011a). Apart from salt tolerance, NHX antiporters are known to involved in flower coloration, K+ homeostasis, cell expansion, vesicular trafficking and protein targeting (Apse et al., 1999; Bowers et al., 2000;
Yamaguchi et al, 2001; Venema et al., 2003; Brett et al., 2005; Ohnishi et al., 2005; Pardo et al.,
2006; Apse and Blumwald, 2007; Hernández et al., 2009; Yoshida et al., 2009; Bassil et al., 2011b;
Leidi et al., 2010). In addition to these, Apse et al. (2003) showed that NHX1 is involved in seedling establishment and leaf development. A study by Bassil et al. (2011b) showed that both
NHX1 and NHX2 helped to control vascular pH, K+ homeostasis, growth, flower development and reproduction in plants. Under salt stress, NHX5 and NHX6 localized in the Golgi and trans-Golgi network are involved in the trafficking of endosomal cargo to the vacuoles and in cell expansion
62 (Bassil et al., 2011a).
HKT transporters are carrier-type proteins which involved in the Na+ and K+ transport in the plasma membranes (Haro et al., 2005). Transporters belonging to the HKT family are classified into two subfamilies (Horie et al., 2009; Yao et al., 2010). HKT transporters belonging to the subfamily 1 have a highly conserved serine residue and preferentially transport Na+ whereas, subfamily 2 have a highly conserved glycine residue and transport both Na+ and K+ in plant cells (Horie et al., 2009;
Yao et al., 2010). Studies pointed out that loss of the function of the HKT1;1 gene in A. thaliana caused accumulation of Na+ in leaves compared to roots (Rus et al., 2004; Sunarpi et al., 2005;
Demidchik and Maathuis, 2007). Further studies on AtHKT1;1 and its rice homolog OsHKT1;5 showed that the Na+ transporter helped to protect photosynthetic tissues by removing excess Na+ from xylem sap into surrounding xylem parenchyma cells (Ren et al., 2005; Sunarpi et al., 2005;
Horie et al., 2006; Davenport et al., 2007). Another study on two rice cultivars, salt-tolerant indica and salt-sensitive japonica pointed out that OsHKT1;4 limited the leaf sheath-to-blade Na+ transport under salinity (Cotsaftis et al., 2012). These findings suggested that HKT subfamily 1 localized on the plasma membrane of xylem parenchyma cells restrict the entry of excess Na+ into leaves and protect photosynthetic tissues (Deinlein et al., 2014). A study on rice HKT showed that, under K+ deprived conditions, OsHKT2;1 catalyzes the uptake of Na+ in roots where Na+ partially replaces the function of K+ and OsHKT2;2 is responsible for the Na+- dependent K+ uptake by roots
(Horie et al., 2007). It was found that OsHKT2;1 is expressed in root epidermis, cortical cells and vascular tissues in roots and leaves (Gollack et al., 2002; Garciadeblás et al., 2003; Horie et al.,
2007). Compartmentalization of Cl- ions through ion channels is an important stress tolerance mechanism in plants (Xu, 1999; Brini and Masmoudi, 2012). Voltage gated ion channels belonging
63 to the CLC (chloride channel) family are known to be involved in the vacuolar Cl- sequestration
(Hechenberger et al., 1996; Barbier-Brygoo et al., 2000; Brini and Masmoudi, 2012). A study of
Diédhiou and Golldack (2006) showed the importance of OsCLCc in the osmotic adjustment of salt treated rice plants. Apart from Cl- homeostasis, these channels are also function as a H+-coupled antiporters and are involved in nitrate accumulation (De Angeli et al., 2006).
Apart from the ion compartmentalization, salt secretion and exclusion also occur in plants as a salt tolerance mechanisms (Parida and Das; 2005; Munns and Tester, 2008). Salt-induced increases in cell size due to a rise of vacuole volume and excretion of Na+ and Cl- ions through salt glands or bladders are the most common anatomical adaptation of halophytes to thrive in saline soil (Flowers et al., 1977).
2) Accumulation of compatible solutes
Accumulation of compatible solutes or osmolytes is another protective mechanism of plants used to survive under salt stress (Hasegawa et al., 2000; Parida and Das, 2005). Compatible solutes including proline, glycine betaine and polyols, help to maintain the osmotic balance without interacting with biochemical reactions in plants (Hasegawa et al., 2000; Parida and Das, 2005; Khan et al., 2000; Bohnert et al., 1995). In addition, sugars, including glucose, fructose, sucrose and fructans accumulate in plants and provide osmoprotection, osmotic adjustment, carbon storage and radical scavenging under salt stress (Parida et al., 2002). Apart from their well known function as osmoprotectants, these solutes are also act as low molecular weight chaperones, involved in stabilizing the PSII complex and protecting the structures of enzymes and proteins (Robinson and
Johnson, 1986; Smirnoff and Cumbes, 1989; McCue and Hanson, 1990; Santoro et al., 1992;
64 Bohnert et al., 1995; Papageorgiou and Murata, 1995; Shen et al., 1997; Hare et al., 1998; Mansour,
1998; Noiraud et al., 2001). High amounts (more than 40 mM on a tissue water basis) of proline/ glycine betaine are found in halophytes compared to glycophytes (Flower et al., 1977). However, in glycophytes, the amount of proline/ glycine betaine is sufficient to generate a significant osmotic pressure in cells under salt stress (Flower et al., 1977). Sickler et al. (2007) showed accumulation of mannitol was enhanced when a mannose-6-phosphate reductase from celery was over-expressed in
Arabidopsis thaliana plants. Although compatible solutes play a major role in stress tolerance in plants, it comes with an energy cost (Munns and Tester, 2008). A study by Raven (1985) showed synthesis of mannitol required 34 ATP, 41 ATP for proline, 50 ATP for glycine betaine and approximately 52 ATP for sucrose.
3) Involvement of the antioxidant enzymes
Salt in soil and water encourages over-reduction of the electron transport chain in mitochondria and chloroplasts, photorespiration, fatty acid oxidation and activity of cell wall peroxidases, germin-like oxalate oxidases and amine oxidases in the apoplast which lead to the production of reactive oxygen species (ROS) (Mittler et al. 2004; Miller et al., 2010). Higher plants have enzymatic and non- enzymatic antioxidant system which scavenge reactive oxygen species (ROS) and other free radicals. The antioxidant system of plants includes low molecular mass non-enzymatic antioxidants, like ascorbic acid, glutathione and tocopherols, and enzymatic antioxidants like SOD (superoxide dismutase), peroxidases and catalases (Nagalakshmi and Prasad, 2001; Shi and Zhu, 2008; Sharma and Dietz, 2009; Ashraf, 2009; Jaleel et al., 2009). Apart from this plants also have phenolic compounds which act as antioxidants, including flavanoids, tanins and lignin precursors (Blokhina et al., 2003). ROS such as H2O2 and superoxides cause damage to plants by peroxidation of unsaturated fatty acids in membranes, desaturation of proteins and disrupting carbohydrates and
65 DNA in cells (Zhang et al., 2001; Parida and Das, 2005; Jithesh et al., 2006).
Antioxidant systems act as a cooperative network when it comes to inhibiting the formation of reactive oxygen and cells contain more than one antioxidant to breakdown ROS. For example, antioxidant enzymes including ascorbate peroxidase and glutathione reductase work together to breakdown H2O2 produced in cells (Suzuki and Mittler, 2006). SOD, which is discovered by
-. McCord and Fridovich in 1969, converts O2 (superoxide) to O2 and H2O2 (Bowler et al., 1992).
SOD has several forms depending on the metal ions in the active site including Cu/Zn SOD,
MnSOD, FeSOD and NiSOD (in Streptomyces) (Kim et al., 1996; Ahmad et al., 2010). Cu/Zn SOD is found to be distributed in the cytosol and chloroplasts and MnSOD is mainly located in mitochondria and peroxisomes. Catalases mainly locate to peroxisomes. Catalases are the principal scavenging enzymes which converts toxic H2O2 to O2 and water (Asada, 1994). Plants have numerous isozyme forms of catalase. These enzymes are categorized into classes (Willekens et al.,
1994; Ahmad et al., 2010). Class 1 catalases are mainly found in photosynthetic tissues and remove
H2O2 produced during photorespiration. Class 2 catalases are localized in vascular tissues and are responsible for the lignification of vascular cells. However, the scavenging role of Class 2 catalases remains unknown. Class 3 catalases are found in seeds and young plant tissues. The biological function of this group is to remove H2O2 during fatty acid degradation in glyoxysomes (Willekens et al., 1994; Ahmad et al., 2010). Ascorbate peroxidase and glutathione reductase are two other scavenging enzymes responsible for breaking down H2O2. Ascorbate peroxidase scavenges H2O2 in the water-water and ascorbate-glutathione cycles (Asada, 1994). Glutathione reductase catalyzes
NADPH-dependent reactions and is localized in the chloroplast stroma, mitochondria, cytosol and peroxisomes (Ahmad et al., 2010). Apart from that, peroxiredoxins (Prx) belong to the enzyme group of peroxidases and play a major role in antioxidant defence systems, to detoxify ROS (Rhee et al., 2001; Dietz, 2011). Prx is also involved in the dithiol-disulfide redox regulatory network of
66 the plant and cyanobacterial cell (Dietz, 2011).
Ascorbic acid (vitamin C) is a widely studied non-enzymatic antioxidant in plants. This antioxidant is found in every plant tissue (Smirnoff, 1996). Ascorbic acid is known to be synthesized in mitochondria and then transported to other cell compartments. Ascorbic acid protects plants from
H2O2 and other toxic free radicals. Apart from scavenging free radicals, ascorbic acid is also involved in the regulation of plant growth, differentiation of cells and metabolism (Smirnoff, 1996).
Ascorbic acid is known to be involved in the regeneration of antioxidant tocopherols (Horemans et al., 2000; Ahmad et al., 2010). Tocopherols have several forms, such as α, β, γ and δ-tocopherols,
Among these, α-tocopherols are the most active form of antioxidant and are widely found in chloroplasts (Munné-Bosch, 2005). α-tocopherols protect plants by quenching singlet excited oxygen by charge transfer mechanisms. Glutathione is a tripeptide and is found in all cells and organelles in plants (Noctor et al., 2012). Glutathione plays a major role in plant protection by being involved in the ascorbate-glutathione cycle. This antioxidant has the ability to scavenge the most dangerous ROS and free radicals including H2O2, singlet excited oxygen, superoxides and hydroxyl radicals. The antioxidant role of carotenoids have been extensively studied and are known to involve quenching singlet excited oxygen in PSII (described in the Section 1.4.1.1. Reactive
Oxygen Species formation). Phenolic compounds in plants including flavonoids, tanins and lignin form polyphenols and act as hydrogen and electron donors. thereby stabilizing unpaired electrons in reactions (Ahmad et al., 2010).
4) Involvement of plant hormones
Salt stress encourages the production of plant hormones (phytohormones) including, abscisic acid
67 (ABA), jasmonates and cytokinins (Thomas et al., 1992; Aldesuquy, 1998; Vaidyanathan et al.,
1999). ABA, considered as the plant stress hormone, acts as an endogenous messenger which induces various stress responsive genes (deBruxelles et al., 1996; Swamy and Smith, 1999). Apart from that, ABA plays a major role in seed dormancy, delays seed germination (encourage seeds to surpass stress conditions and germinate only when the conditions are favorable), development of seeds, stomatal closure, embryo morphogenesis, synthesis of storage proteins and lipids, leaf senescence and defense against pathogens (Swamy and Smith, 1999). A study by Thomas et al.
(1992) showed that in Mesembryanthemum crystallinum, ABA induces a switch from C3 photosynthesis to crassulacean acid metabolism (CAM) under salinity. One of the main functions of
ABA in the stress tolerance is regulating the water and osmotic balance under water deficit conditions (Tuteja, 2007). Koornneef et al. (1998) showed that although the growth rate of ABA mutants of A. thaliana, aba1, aba2 and aba3 is similar to the wild type, they readily wilted and died if drought persisted. Stress-induced plant responses can be ABA-dependent, ABA-independent or partially ABA-dependent (Zhu, 2002). Most ABA inducible genes share regulatory elements called
ABA-responsive element (ABRE) (Thomashow et al., 1999; Shinozaki and Yamaguchi-Shinozaki,
2000; Uno et al., 2000; Zhu, 2002). Although there are two pathways, genetic analysis indicated that there is no clear separation between these, which are interconnected through calcium (Swamy and
Smith, 1999; Xiong et al., 2002; Chinnusamy et al., 2004; Mahajan and Tuteja, 2005). A study by
Knight et al. (1997) pointed out the involvement of calcium in the ABA-dependent induction of
P5CS gene (encodes Δl-pyrroline-5-carboxylate synthetase, P5CS, the first enzyme of the proline biosynthetic pathway) in A. thaliana under salt stress. Other than that, proteomic studies have shown several ABA-related proteins including, ABA-responsive proteins (ABR17 and ABR18) and
ABA or salt-induced protein (ASR1) in Pisum sativum and Oryza sativa under salt stress (Salekdeh et al., 2002; Kav et al., 2004). These proteins are associated with the salt stress tolerance in many plants (Tuteja, 2007). A study from Srivastava et al. (2006) showed that over-expression of pea
68 ABR17 in A. thaliana induced the expression of proteins like DNA damage repair proteins and photosynthetic proteins under salt stress.
Other than ABA, jasmonates (methyl jasmonate, MeJA and jasmonic acid, JA) are important phytohormones involved in stress regulation in plants (Cheong and Choi, 2003). Major functions of
JA are root growth, seed germination, fertility, fruit ripening and leaf senescence (Wasternack and
Hause, 2002). A study by Pedranzani et al. (2003) pointed out that in tomato, salt-tolerant cultivars showed high levels of jasmonate compared to the salt-sensitive cultivars. Kang et al. (2005) found an increased level of JA in salt-tolerant rice cultivars compared to salt-sensitive cultivars. Similarly another study showed a significant rise in the levels of MeJA in rice roots when exposed to 200 mM of NaCl (Moons et al., 1997). According to these studies, it is evident that the concentration of endogenous JA increases as a protective mechanism against salt stress. However, very little information is available on the factors involved in the regulatory mechanism of endogenous JA
(Parida and Das, 2005; Javid et al., 2011). Studies on the effects of exogenous JA showed that application of JA after salt treatments enhance the salt tolerance in plants (Tsonev et al., 1998; Kang et al., 2005). This suggested that exogenous JA changes the balance of other stress response hormones such as ABA, which promote stress tolerance mechanisms in salt stressed plants (Kang et al., 2005; Javid et al., 2011).
1.3.1.5. Effects of salt stress on other photosynthetic organisms
1) Cyanobacteria
Cyanobacteria are oxygenic phototrophic bacteria and the only prokaryotes in their photosynthesis similar to plants. Many proteomic and transcriptomic studies on stress tolerance have been
69 performed using cyanobacteria (Pandhal et al., 2008a). Given their similarity to chloroplasts, cyanobacteria act as an ideal model to identify plant cellular functions and metabolisms under salt stress (Pandhal et al., 2008a; Pandhal et al., 2008b). Cyanobacteria are the only living organisms which are capable of both photosynthesis and the biological nitrogen fixation (Gray and Doolittle,
1982). They colonize a wide range of ecosystems such as soil and aquatic systems. They can categorize into three groups: salt sensitive (stenohaline), moderately halotolerant and extremely halotolerant (Reed and Stewart, 1988; Pandhal et al., 2008b). Among cyanobacteria, Synechocystis sp. PCC6803 is a unicellular, fresh water, oxygenic and moderately halotolerant cyanobacteria, which is extensively used to understand the physiology and the biochemistry of salt stress tolerance
(Pandhal et al., 2008a). The genome of Synechocystis sp. PCC6803 was sequenced in 1996 and this strain can be easily use for genetic manipulation (Grigorieva and Shestakov, 1982; Kaneko et al.,
1996).
In Anabaena torulosa transcription of almost 10% of entire genome is been regulated by changes in the salt concentration (Apte and Haselkorn, 1990). Microarray analysis has shown that salinity induces many genes involved in antioxidant defence system including genes for heat shock proteins
(hspA, dnaK, dnaJ, htrA, groEL2, clpB) and superoxide dismutase (sodB) (Campbell and
Laudenbach, 1995; Lee et al., 1998; Roy et al., 1999; Nakamoto et al., 2000). Apart from that, genes such as glpD and ggpS involved in the synthesis of osmoprotectant, glucosylglycerol (GG), genes coding for proteases (HtyA, ClpB) and chaperone proteins (DnaK, DnaJ, GroEL) were found through microarray analysis (Kanesaki et al., 2002; Marin et al., 2003; Marin et al., 2004;
Shoumskaya et al., 2005; Castielli et al., 2009). Additionally, using the subtractive RNA hybridization procedures, several other genes, including cpn60 (encoding GroEL), isiA
(chlorophyll-binding protein), crh (RNA helicase) and petH (ferredoxin: NADP1) were identified as being induced in Synechocystis sp. PCC6803 under salt stress (Vinnemeier and Hagemann, 1999).
70 This cyanobacterial strain is also used to identify the interaction between hik (Histidine kinase) and
Rre (response regulator). This two-component system is involved in the perception and signal transduction pathway when cells are exposed to low temperature, salt stress, osmotic stress and metal ion deficiencies (Suzuki et al., 2001; Yamaguchi et al., 2002; Karandashova et al., 2002;
Marin et al., 2003; Paithoonrangsarid et al., 2004). It was found that under stress conditions, hik phosphorylated and activated Rre, which is responsible for the expression of genes involved in stress regulation (Marin et al., 2003; Paithoonrangsarid et al., 2004). According to microarray analysis, five hik-Rre systems exist in Synechocystis sp. PCC6803 under salt stress (Shoumskaya et al., 2005).
Excess salt in living cells causes imbalances in K+, which is important for cellular homoestasis by maintaining cell turgor (Alahari and Apte, 1998). H+/Na+ and Na+/K+ antiporters are involved in Na+ efflux from cells. This is driven by proton gradients across membranes with the involvement of cytochrome oxidases and/or by hydrolysis of ATP. Kanamaru et al. (1994) showed that the P-type
ATPase in the thylakoid membranes of Synechococcus sp. PCC7942 maintained the ion homoestasis in cells by active extrusion of Na+. Another study from Apte and Thomas (1986) showed that aa3-type cytochrome oxidase in plasma membranes in two Anabaena species involved in the excess Na+ extrusion from cells. Although, the salt tolerance mechanisms in cyanobacteria vary according to the species, the basic mechanism of Na+ transport appears to be identical (Apte and Thomas, 1986). Apart from this, cyanobacteria also have Ca2+-ATPases and ion channels for
Ca2+ accumulation which is important for K+/Na+ selectivity of cells. Work from Raeymaekers et al.
(2002) showed that Bacillus subtilis consists of P-type Ca2+ transport ATPase during sporulation.
Apart from ion channels and ATPase, membrane fluidity plays a major role in cell adaptation to excess salt (Singh et al., 2002). A study by Allakhverdiev et al. (2001) showed that in
Synechococcus sp. the unsaturation of the fatty acids in membrane lipids associated with the
71 photosynthetic apparatus increases salt tolerance.
The effects of salt on photosynthetic pigments varies according to the type of the pigment. For example salinity decreases chlorophyll content in Anabaena doliolum as well as phycobiliproteins
(Singh and Kshatriya, 2002; Srivastava et al., 2005). However, carotenoids content is enhanced under salt stress and provides protection to chlorophyll by acting as an antioxidant (Srivastava et al.,
2005; Srivastava et al., 2006). A study performed by Sudhir et al. (2005) showed a decline in PSII photochemistry and increase in the PSI activity in Spirulina platensis. Degradation of D1 protein is suggested as the main reason for the decline in PSII activity (Ohad et al., 1984; Rintamak et al.,
1994; Sudhir et al., 2005). The increase in the PSI activity is suggested to be the activation of cyclic phosphorylation and this was observed in several cyanobacteria species including Spirulina platensis and Synechocystis PCC6803 (Schubert and Hagemann, 1990; Jeanjean et al., 1993; Sudhir et al., 2005).
Cyanobacteria also accumulate non-toxic organic compounds which act as osmoprotectants, including sucrose, trehalose, glucosylglycerol (GG), glutamate and glycine betaine under salt stress
(Reed et al., 1985; Welsh et al., 1996; Marin et al., 1998; Marin et al., 2006; Bhargava et al., 2008;
Yoshikawa et al., 2011; Reina-Bueno et al., 2012). Among those, GG is the extensively studied osmoprotectant in cyanobacteria (Srivastava et al., 2011). Work from Mikkat et al. (1996) showed a system which involved in the transportation of GG in Synechocystis PCC6803. In addition to that a study of Csonka and Hanson (1991) described a pathway of glycine betaine in cyanobacterium,
Aphanothece halophytica.
72 2) Algae
Algae are important organisms as they are inhabitants of many biotopes with changing salinities
(Bohnert and Jensen, 1996; Bohnert and Sheveleva, 1998; Fogg, 2001). Therefore, they serve as model organisms for a better understanding of the mechanisms of salt stress regulation (Bohnert and
Jensen, 1996; Bohnert and Sheveleva, 1998; Fogg, 2001). Among alga, Chlamydomonas reinhardtii is the most widely studied and used laboratory strain (Mastrobuoni et al., 2012). Neelam and
Subramanyam (2013) showed that 150 mM of salt caused a reduction in cell size, flagellar resorption, slower growth rates, reduction in chlorophyll pigments and clumped morphology in C. reinhardtii. The reduction of electron transport to reaction centers may have occurred due to the damage of core proteins CP43 and CP47 in the PSII complex (Neelam and Subramanyam, 2013).
Studies have shown that hyperosmotic stress caused by excess Na+ and Cl- in cells, damages the structure of PSI-LHCI (Subramanyam et al., 2010) and inhibits electron transport between plastocyanin and P700+ (Cruz et al., 2001) in C. reinhardtii. Marín-Navarro and Moreno (2006) showed that salinity caused oxidation and degradation in Rubisco enzymes in C. reinhardtii.
Similar to plants, salinity induces several signalling pathways in algae (Arisz and Munnik, 2011).
Sudden increases in phosphatidic acid and lysophosphatidic acid concentrations in Chlamydomonas imply that salinity induces phospholipid signalling pathways (Meijer et al., 2001; Meijer et al.,
2002; Arisz et al., 2003; Arisz and Munnik, 2011). 76 salt induced proteins were identified in halotolerant green algae, Dunaliella salina (Liska et al., 2004). In this proteomic study, nanoelectrospray mass spectroscopy was combined with sequence similarity database searching algorithms, MS BLAST and multiTag to identify 80% of salt induced proteins associated with the
Calvin cycle, starch metabolism, redox energy production, regulatory factors in protein synthesis and degradation. These suggested that this halotolerant algae is capable of enhancing
73 photosynthesis under salinity by increasing the CO2 assimilation and by diversion of carbon and energy source for the synthesis of glycerol (Liska et al., 2004). Proteomic and metabolomic studies performed by Mastrobuoni et al. (2012) showed that salt has major impacts on the amino acid metabolism and induces proline biosynthesis in C. reinhardtii. A comparative proteomic analysis was performed using C. reinhardtii after exposing them to 300 mM of salt for a short period of time
(2 hours) (Yokthongwattana et al., 2012). In this study, they showed that a number of proteins were exclusively found in one sample but not the other. 18 proteins were only found in control samples and they are mostly involved in general metabolic pathways. 99 proteins uniquely appeared in salt- treated sample, which are mostly stress related proteins and involved in protein translation such as eukaryotic initiation factor EIF31 and elongation factor Tu. This study and the previous proteomic study performed on Dunaliella salina (Liska et al., 2004) suggested that most salt induced genes encode proteins which are involved in the translation machinery.
Expressed sequence tag (EST) analysis were done to identify salt responsive genes in the red alga
Furcellaria lumbricalis (Kostamo et al., 2011). Transcriptomic analysis were performed to understand the short-term (after 48 hours) and long-term (after 1255 generations) acclimation of C. reinhardtii exposed to 200 mM of NaCl (Perrineau et al., 2014). In this study, as the short-term responses they found that cells exhibit many well-known stress responses including low photosynthetic rate, upregulation of salt-responsive genes, upregulation of glycerophospholipid signalling and transcription and translation machinery. However, after 1255 generations, cells exhibit downregulation of stress responsive genes and down regulation of glycerophospholipid signalling. These findings suggested that long term exposure enhances the adaptation of C. reinhardtii to salt.
74 1.3.2. Drought stress
Water is essential for living organisms and the major medium for transporting important metabolites and nutrients (Hsiao, 1973). Water deficit lowers the plant water potential and turgor. Therefore, plants lack the ability to perform almost all the physiological and biological functions (Tuba et al.,
1996; Sarafis, 1998; Yordanov et al., 2003; Zlatev and Lidon, 2012). Water stress of plants occurs when soils lack water which can be absorbed by roots or when the transpiration rate of leaves is higher than the water absorption by roots. Accumulation of solutes in the cytosol occurs due to the loss of water. Lack of water inside the plant cells results in shrinking of the cell, which restrict plant growth and reproduction (Tuba et al., 1996; Sarafis, 1998; Yordanov et al., 2003; Zlatev and Lidon,
2012). Apart from affecting plant-water relations, drought also causes adverse effects on other physiological processes, including stomatal closure, restricting gas exchange, reducing transpiration and inhibiting photosynthesis (Cornic, 1994). Photosynthesis is severely restricted by drought. Low photosynthetic rates initially occur due to the closure of stomata and then, as drought becomes more severe, by metabolic impairment under water deficient conditions (Cornic et al., 1989; Sharkey,
1990; Cornic and Briantais, 1991; Giménez et al., 1992; Tezara and Lawlor, 1995; Panković et al.,1999).
C4 plants have higher water-use efficiency than C3 plants. Therefore, C4 photosynthesis is considered as less sensitive to drought than C3 (Haxeltine and Prentice, 1996; Taub, 2000; Cabido et al., 2007). However, several studies have shown that C4 plants become more sensitive to drought under severe water deficit conditions (Ellis et al., 1980; Medrano et al., 2002; Flexas et al., 2004;
Ripley et al., 2007; Osborne, 2008; Ripley et al., 2010). Drought affects the chlorophyll content of plants by inhibiting chlorophyll synthesis (Lisar et al., 2012). Unlike chlorophylls, carotenes and xanthophylls are less sensitive to drought (Niyogi et al., 1997). Drought has an adverse effect on the
75 Studies have revealed that decreased photosynthetic capacity results from impaired regeneration of ribulose‐‐ 1,5 bisphosphate (RuBP) (Giménez et al., 1992). Studies have suggested that this may be due to decreased ATP synthesis (Gunasekara and Berkowitz, 1993; Tezara et al., 1999). Apart from that, water deficit conditions decrease the level and the activity of Rubisco (Tezara et al., 1999).
Although the Rubisco holoenzyme is relatively stable under drought stress (Webber et al., 1994), studies have shown that drought leads to a rapid decrease in the abundance of Rubisco small subunit
(rbcS) transcripts in tomato (Bartholomew et al., 1991), A. thaliana (Williams et al., 1994) and rice
(Vu et al., 1999). A study by Parry et al. (2002) suggests that in tobacco the decrease of Rubisco
2+ activity under drought stress is not primarily the result of changes in activation by CO2 and Mg but due rather to the presence of tight‐ binding inhibitors.
Mineral uptake of plants is severely affected by drought. Tomato plants showed a reduced level of nitrogen and phosphorous under drought stress (Subramanian et al., 2006). Similarly, Asrar and
Elhindi (2011) showed a reduction in phosphorous content in marigold seedlings under water deficit conditions. Drought causes a major change in protein and lipid content in plants. Although proteins in plant leaves decrease due to water deficit conditions, stress induced proteins including Hsps
(heat-shocking proteins) and LEAs (late embryogenesis abundant) are increased (Al-Whaibi, 2011;
Hand et al., 2011; Lisar et al., 2012). Plants undergo anatomical, morphological and cytological changes to withstand drought conditions, including leaf size reduction, decreases in the number of stomata, thickening of cell walls and cutinization of leaf surfaces.
1.3.3. Heat stress
Temperature fluctuations throughout the world affect the growth and development of plants. Global
76 warming is considered as a major reason behind environmental and ecological changes, due to the temperature increases (Loik et al., 2000; Kipp, 2007). It has been found that the global mean temperature has increased by 0.6 oC over past 100 years and according to global temperature models, the temperature will increase by 2-6 oC over next 100 years (Kipp, 2007). Therefore, it is necessary to find methods to produce plants able to withstand the rising temperatures. Heat stress is known to cause adverse effects on plant growth and reproduction (Havaux and Davaud, 1994;
Gulen and Eris, 2004). High temperatures decrease the size of the A. thaliana plants and accelerate flower development compared to control plants (Kipp, 2007).
Loik and co-workers (2000) found a decreased quantum efficiency of PSII and increased NPQ in an evergreen shrub (Artemisia tridentata) and a herbaceous forb (Erigeron speciosus) when exposed to heat. Some plants show decreases in anther and pollen development under heat stress. A study with tomato showed an impairment of pollen and anther development under increased temperature and decreased fruit set (Peet et al., 1998). Similarly, common beans showed an abnormal pollen and anther development during microsporogenesis under elevated temperature (Porch and Jahn, 2001).
Another study on flax plants (Linum usitatissimum L.) showed decreases in flowering, seed set, pollen viability and germinability under heat stress (Cross et al., 2003).
1.3.4. Low temperature stress
Low temperatures can result in poor growth and development in plants all over the world, especially in tropical and sub-tropical regions. Sensitive plants show noticeable physiological dysfunction at temperatures under 10 to 12 oC. This is considered as chilling injury (Lyons, 1973). Temperatures below zero cause freezing injury to plants. Barthel et al. (2014) showed that sudden decrease in
77 temperature (from 25 to 10 oC) delays plant carbon transport and invest relatively more carbon into respiration than growth or storage. Tolerance levels of plants for low temperatures depends on the region of origin (Lyons, 1973). For example, temperate plants including apple, can cope with temperatures around 0 to 2 oC, sun-tropical plants including pineapple and citrus, can only survive in temperature around 8 oC and tropical plants including banana can only cope temperatures around
12 oC (Lyons, 1973).
Plants show a reduction in growth and development under low temperatures. Ercoli and co-workers
(2004) pointed out that sorghum plants exposed to low temperatures (2, 5 and 8 oC) showed decreased growth and low nitrogen uptake. They also discussed that sorghum plants were able to harden to low temperature when exposed for a long time. However, this ability drops with decreasing temperature. Apart from growth inhibition, low temperatures affect fruits (Adams et al.,
2001). The most prominent chilling injuries on fruits are surface pitting, necrotic areas and external discolouration (Wang, 1994). Cucumber is considered as highly sensitive to chilling temperatures.
Surface pitting is one of the common symptoms occurring in cucumber when exposed to low temperatures (Lyons, 1973). Chilling temperatures cause adverse effect on photosynthesis of plants.
Oxidative damage occurs in electron transport chain, due to the over-excitation of the reaction centres and carbon reduction is inhibited due to decreases in the activity of Rubisco and stomatal closure (Allen and Ort, 2001).
Freezing temperatures or sub-zero temperatures cause adverse effects on plant productivity and limit the distribution of plants (Thomashow, 1998). When the temperature drops below zero, ice can form in the intercellular spaces in plant tissues. This causes plant cells to suffer from dehydration, due to the flow of unfrozen water from inside the cell to the outside. This leads to denaturation of cellular proteins and precipitation of various molecules (Thomashow, 1998). Plants exhibit several
78 adaptations to survive in extreme low temperatures. Plants show a xerophytic nature to survive in the low water conditions. Some plants have high intracellular solute concentrations and encourage ice nucleation outside the cells (Thomashow, 1998; Allen and Ort, 2001). Plants activate reactive oxygen scavenging enzymes to protect plants from ROS due to cold stress (Schöner and Krause,
1990; Prasad, 1997). Changes in the lipid composition of membranes is a common adaptation to cold stress. Plant membrane lipids change from a gel to a liquid-crystalline phase through lipid desaturation under low temperatures (Lyons, 1973; Allen and Ort, 2001).
79 1.4. Effects of environmental stress on the electron transport of photosynthesis
Photosynthesis is one of the main physiological processes responsible for plant growth (Munns et al., 2006; Chaves et al., 2009). Abiotic stresses such as drought, high light and salinity affect photosynthesis of plants and cyanobacteria (Sudhir et al., 2005). However, plants show complex photosynthetic responses to abiotic stress. This section aims to introduce some of the effects of environmental stress on photosynthesis. However, it only focuses on discussing the effects which are more relevant to this thesis. Firstly, a detailed discussion is given about the impact of environmental stress on energy use in photosynthesis and how inhibition occurs. In the same section the production of reactive oxygen species (ROS) under stress conditions is discussed. Then a detailed discussion is given on the effects of abiotic stresses on protein components of the electron transport.
1.4.1. Effects on energy use in photosynthesis
Light absorbed by chlorophyll-a molecules converts it into its singlet excited state (1Chl*). 1Chl* returns to the ground state by releasing energy in one of several ways: as chlorophyll fluorescence, by transferring energy to reaction centres to drive photosynthesis or as heat through non- photochemical quenching (NPQ) (Müller et al., 2001). The energy in 1Chl* can also be dissipated through converting 1Chl* to triplet excited state (3Chl*) (Foyer and Harbinson, 1999). 3Chl* has the potential to transfer energy and produce singlet excited oxygen which is extremely toxic to plants
(Müller et al., 2001).
80 Three processes; photochemistry, heat and fluorescence are in competition with each other and an increase in one will result in a decrease in the yield of other two (Maxwell and Johnson, 2000).
Fluorescence is measured as the fluorescence spectrum, which is different to that of the absorbed light and the peak of fluorescence emission has a longer wavelength than that of absorption
(Maxwell and Johnson, 2000). Chlorophyll fluorescence analysis is an extensively used technique to study the energy use efficiency of PSII in plants.
The term NPQ describes all processes that lower the yield of chlorophyll fluorescence by dissipating energy as heat (Maxwell and Johnson, 2000; Müller et al., 2001). There are three major processes that contribute to NPQ and these can be partly distinguished by their relaxation kinetics.
First is high energy state or pH dependent quenching (qE) (Horton et al., 1996). This is considered as most important component of NPQ and is found in both plants and algae. qE relaxes within seconds to minutes (discussed in Section 1.5.2). The second component is qT, state-transitions quenching important in particular for algae. qT relaxes within tens of minutes (Walters and Horton,
1991) (discussed in Section 1.5.1). The third component is photoinhibition quenching (qI)
(Osmond, 1994) (discussed in Section 1.4.2). This quenching is caused by the photoinhibition of
PSII and relaxes over long period of time (hours) (Maxwell and Johnson, 2000; Müller et al., 2001).
Nilkens et al. (2010) introduced a new component of NPQ: zeaxanthin dependent component (qZ) in A. thaliana. According to this study, the formation (lifetime 10-15 minutes) and relaxation of this component correlated with the synthesis and epoxidation of zeaxanthin. However, comparative analysis of different mutants of A. thaliana suggested that qZ does not show any relationship with other components (qE, qT, qI) of NPQ and therefore represent as a separate component of NPQ.
It is essential that the energy and reducing power produced by photosynthetic electron transport are kept in balance with the requirements of CO2 fixation to prevent ROS production (Cruz de
81 Carvalho, 2008; Miller et al., 2010; Foyer and Shigeoka, 2011). Abiotic stresses like salinity and drought cause stomatal closure to protect from excess transpiration. Closure of stomata disturbs gas exchange in leaves. This causes reductions in photosynthesis. When CO2 fixation is inhibited, the electron transport chain become over excited with excess light, which encourages the production of reactive oxygen species (Smirnoff, 1993; Golding and Johnson, 2003; Chaves et al., 2009; Saibo et al., 2009; Stepien and Johnson, 2009). Molecular oxygen is considered as relatively inactive.
However, molecular oxygen is subsequently converted into more reactive forms such as superoxide, hydrogen peroxide, hydroxyl radicals and singlet oxygen. These molecules are potentially dangerous to cellular components including membranes, proteins and DNA (Shah et al., 2001;
Mittler, 2002; Verma and Dubey, 2003; Meriga et al., 2004; Sharma and Dubey, 2005; Maheshwari and Dubey, 2009; Mishra et al., 2011; Srivastava and Dubey, 2011).
1.4.1.1 Reactive oxygen species (ROS) formation
The reaction centres of photosystems I and II are the main sites for reactive oxygen formation. In
1951, Mehler discovered the photoreduction of O2 to H2O2 in PSI. The primary reduced product was
. - identified as superoxide ( O2 ) which is then disproportionated to produce O2 and H2O2 (Asada,
2006). In PSII, energy absorbed by P680 is converted to a triplet excited state, 3P680* (results from charge recombination within the radical pair P680+Pheo- when PSII acceptors are reduced) (Hideg et al., 1994; Hideg et al., 1998; Ivanov and Khorobrykh, 2003). Then energy is transferred to
3 1 oxygen ground state (triplet, O2) to form excited singlet oxygen ( O2*) (Equation 1) (Telfer et al.,
1994; Asada, 2006).
Equation 1:
3 3 1 P680* + O2 O2* + P680
82 1 Singlet excited oxygen ( O2*) is highly reactive and interacts with the molecules in its immediate
1 surroundings (Krieger-Liszkay, 2005). O2* causes damage to the D1 protein in the reaction centres of PSII and disturbs electron transport (Vass et al., 1992). Apart from that, singlet excited oxygen can form other highly active oxygen free radicals. It is mainly produced at the acceptor side of PSII.
Over-reduction of QA, which is a one-electron acceptor, due excess light, causes charge
3 1 recombination from Pheo to oxidized P680 to form P680* and O2* (Vass et al., 1992; Yamamoto et al., 2008). The donor side of PSII is known to produce cationic free radicals including P680+ and
+ Tyrz (Tyrz is the secondary donor of PSII). Both types of ROS cause damage to PSII (Yamamoto et al., 2008). In mild stress conditions, plants manage to maintain the excited oxygen at low levels through the activity of antioxidants and carotenoids. Plants also are able to repair PSII by replacing
D1 protein which is damaged by ROS. However, under severe stress conditions, antioxidant systems and repair mechanisms are unable to prevent oxidative damage and subsequent photoinhibition (Telfer et al., 1994; Asada, 2000).
Apart from capturing light energy in photosystems, carotenoids also play a major role in photoprotection by preventing singlet oxygen formation. Carotenoids are able to protect plants by quenching the energy from the triple excited state, 3P680*, thereby preventing the production of singlet oxygen, the energy being dissipated as heat (Equation 2). In addition, carotenoids have the potential to quench energy directly from singlet oxygen and dissipate is as heat (Equation 3)
(Cerullo et al., 2002; Krieger-Liszkay, 2005). α-tocopherol also acts as a scavenger in PSII and quenches energy from singlet oxygen (Kruk et al., 2005).
Equation 2:
3P680* + carotenoid P680 + 3carotenoid* P680 + carotenoid + heat
83 Equation 3:
1 3 O2* + carotenoid O2 + carotenoid* O2 + carotenoid + heat
. - The major site of superoxide ( O2 ) anion production is the FeS centres on the accepter side of PSI.
Superoxide is considered as an intermediate product of the water-water cycle or the Mehler reaction, associated with PSI. Electrons from PSII are donated to molecular oxygen (O2) to form
. - superoxide ( O2 ) anions (Equations 4 and 5). Superoxide is further converted to hydrogen peroxide
(H2O2) and O2. This reaction is catalysed by superoxide dismutase (SOD) (Equation 6) (Asada,
2006).
Equation 4:
- + 2H2O 4e + O2 + 4H (PSII)
Equation 5:
- . - 2O2 + 2e O2 (PSI)
Equation 6:
. - + 2 O2 + 2H H2O2 + O2 (catalysed by SOD)
. . - Hydroxyl radicals ( OH) are generated from hydrogen peroxide (H2O2) and superoxide ( O2 ). This reaction is known as Haber-Weiss reaction (Equation 7) and consists of several steps (Pospíšil et al.,
2004). The production of hydroxyl radicals is catalysed by metal ions like Fe+2 and Zn+2. The second catalytic reaction is known as the Fenton reaction (Step 2). Hydroxyl radicals are known to affect and deactivate Calvin cycle enzymes (Kaiser, 1979).
84 Equation 7:
3+ . - 2+ Step 1) Fe + O2 Fe + O2
2+ 3+ - . Step 2) Fe + H2O2 Fe + OH + OH
. - . - (Net reaction) O2 + H2O2 OH + OH + O2
1.4.2. Effects on components of electron transport
Exposure to high light under abiotic stress conditions encourages the deactivation of PSII. This process in known as photoinhibition (Powles, 1984; Demmig-Adams and Adams, 1992; Melis,
1999; Murata et al., 2007). Photoinhibition is unavoidable in plants under continuous stress conditions. However, the degree of photoinhibition depends on the balance between degradation of
PSII and the repair mechanisms (Murata et al., 2007). PSII is one of the major sites producing ROS under stress. Damage to the D1 protein in the reaction centre of PSII by ROS is the major cause of photoinhibition (Takahashi and Murata, 2008). Photodamge to PSII is an important reaction occurring in all oxygenic photosynthetic organisms (Takahashi and Murata, 2008). Photosynthetic organisms have developed repair mechanisms to avoid the accumulation of damaged PSII. The repair mechanisms of PSII consist of several steps, including proteolytic degradation of the D1 protein using FtsH and Deg proteases (Nixon et al., 2010), synthesis of the precursor to the D1 protein (pre-D1), insertion of the newly synthesized precursor into the thylakoid membrane with the assembly of other PSII proteins, maturation of the D1 protein by C-terminal processing of pre-D1; and assembly of the oxygen-evolving machinery (Aro et al., 1993; Aro et al., 2005; Takahashi and
Murata, 2008). Studies have shown that plants enhance the recovery process of D1 protein under light and the process is delayed in complete darkness. It is also clear that recovery depends on the
85 temperature (Gombos et al., 1994; Yamamoto et al., 2008). Although photoinhibition is a result of the balance between the rate of damage to PSII and the rate repair, prolonged severe stress conditions cause irreversible inhibition of PSII (Takahashi and Murata, 2008). Nishiyama and co- workers (2001) indicated that oxidative stress or ROS stimulate photoinhibition by inhibiting the protein repair mechanism rather than by stimulating photodamage.
There are two mechanisms proposed for the photoinhibition of PSII: acceptor-side and donor-side photoinhibition (Barber and Andersson, 1992; Aro et al., 1993; Yamamoto, 2001). Singlet excited
1 oxygen ( O2*) formed is thought to be responsible for the photodamage of PSII in acceptor-side photoinhibition (Takahama and Nishimura, 1975, Macpherson et al., 1993; Hideg et al., 1994;
Mishra et al., 1994; Telfer et al., 1994; Barber, 1998). Donor side photoinhibition involves cationic
+ + radicals like P680 and Tyrz and cause damage to D1 protein (Jegerschöld et al., 1990; Blubaugh et al., 1991). Other than these two mechanisms of photoinhibition, other studies have suggested the involvement of Mn in photoinhibition of PSII (Hakala et al., 2005; Ohnishi et al., 2005). Jung and
Kim (1990) suggested that singlet oxygen generated in the iron-sulphur centers or cytochrome chromophores is involved in the photoinhibition.
In addition to photodamage in PSII, PSI also shows photoinactivation under stress conditions.
Winter rye, under low temperature and high light showed photoinhibition of PSI due to a low maximum quantum yield in electron transport in PSII, reduced levels of photoxidizable reaction centre pigments and low efficiency of P700+ oxidation (Ivanov et al., 1998). Photoinhibition in PSI is considered as more dangerous, due to slow recovery mechanisms. Photoinhibition of PSI was first discovered in cucumber leaves under chilling temperatures (Terashima et al., 1994). Tropical and sub-tropical plants including cucumber, coffee, cotton and common beans have a high tendency
86 to undergo photoinhibition in PSI when exposed to chilling temperatures (Nakano et al., 2010;
Ramalho et al., 1999; Sonoike and Terashima, 1994; Sonoike, 2011). However, some chilling tolerant plants, like potatoes and A. thaliana also show photoinhibition of PSI (Havaux and Davaud,
1994; Zhang and Scheller, 2004). It has been found out that hydroxyl radicals (OH.-) produced in
PSI disrupt the iron-sulfur centres (Sonoike et al., 1997). Degradation of reaction centre subunits and fragmentation of PsaB, PsaA and other small subunits occur due to ROS in PSI (Ivanov et al.,
1998). Isolated PSI reaction centres show photoinhibition under high light intensities. However, this effect is not observed in intact leaves (Sonoike, 2011).
The recovery of PSI from photoinhibition is slower than that of PSII. In cucumber leaves, photoxidizable P700+ was not fully recovered a week after a photoinhibitory treatment. Most PSI complexes are not repaired but degrade after photodamage (Kudoh and Sonoike, 2002). When considering the relationship between each photosystem in photoinhibition, it is clear that electron flow from PSII is essential for the photoinhibition of PSI. It was found that photoinhibition of PSII prevented the photoinhibition of PSI in vivo (Kudoh and Sonoike, 2002; Sonoike, 2011). However, photoinhibition of PSI encouraged photoinhibition of PSII and decreased repair of D1 protein by inhibiting ATP synthesis (Kudoh and Sonoike, 2002; Sonoike, 2011). A recent study of Tikkanen et al. (2014) showed that, regulation of PSII photoinhibition is the ultimate regulator of the photosynthetic electron transfer chain which provides a photoprotection against reactive oxygen species formation and photodamage in PSI.
Work from Sonoike (1995) pointed out that in isolated thylakoids from cucumber and spinach, photoinhibition of PSI occurred under very low light intensities but this was not observed in vivo.
Therefore, it is clear that photoprotective mechanisms are lost during the isolation of the thylakoids.
87 However, the photoprotective mechanism involved is not yet identified. Cyclic electron transfer from PSI to the plastoquinol pool is considered as a protective mechanism which reduces the over- reduction of PSI and hence protects PSI from damage. Scavenging enzymes, superoxide dismutase
(SOD) and ascorbate peroxidase play a major role in protecting PSI from superoxide anion radicals produced from the reduction of oxygen (Sonoike, 2011). A recent study by Kono et al. (2014) changing light conditions caused stress in PSI and cyclic electron transport around PSI is essential to protect PSI from photoinhibition.
88 1.5. Regulation of electron transport chain of photosynthesis under stress conditions
Being sessile in an ever changing environment requires plants to be physiologically dynamic to survive. Photosynthesis is one of the major metabolic processes which is highly susceptible to abiotic stress. Therefore, plants have regulatory mechanisms to protect the components of photosynthesis from stress. This section focuses on outlining the regulation of photosynthesis of plants under abiotic stress conditions.
1.5.1. State-transitions (qT)
Different protein and pigment compositions cause different light absorption in light harvesting complexes (LHC) in two photosystems (Bonaventura and Myers, 1969; Murata, 1969; Allen, 1992;
Rochaix, 2011). Therefore, photosynthetic organisms need to arrange LHC rapidly, to adjust the relative absorption cross-section of photosystems (Ünlü et al., 2014). The state-transitions, or state
1- state 2 transition, is a mechanism where the light harvesting apparatus of the two photosystems in photosynthesis is remodelled according to changes in light conditions (Allen, 1992). State- transitions are consider as a short-term response, which help plants to protect photosystems from over-excitation and photoinhibition (Wollman, 2001; Lemeille and Rochaix, 2010). This mechanism was first discovered in red algae and green algae over forty years ago (Bonaventura and Myers,
1969; Murata, 1969).
It was discovered that the activation and the de-activation of protein kinases and phosphatases in thylakoid membranes which are regulated through the redox states of the PQ pool and the Fd/TRX
89 systems are essential for state-transitions in photosystems (Allen and Forsberg, 2001; Wollman,
2001; Lemeille et al., 2009). Over-reduction of inter-system electron carriers in electron transport activate a protein kinase and the activated kinase then phosphorylates LHCII subunits (Wollman,
2001). Depending on whether PSII or PSI is preferentially excited, the PQ pool becomes reduced or oxidized under changing light. When the PQ pool is reduced, PQH2 binds to the Qo site of the Cyt b6f, which leads to the activation of a protein kinase that phosphorylates LHCII. After this, the mobile part of LHCII is displaced from PSII to PSI (State 2) (Vener et al., 1997; Zito et al., 1999;
Rochaix, 2013). This process is reversible as overexcitation of PSI cause oxidation of the PQ pool, inactivates the LHCII kinase and results in net dephosphorylation of LHCII by a phosphatase, which returns to PSII (State 1) (Rochaix, 2011; Rochaix, 2013). In C. reinhardtii, state transitions are important, as they play a major role in ATP homeostasis and 80% of the LHCII antennae is mobile, while in plants, only 15-20% can be phosphorylated (Delosme et al., 1996; Vener et al.,
1997).
Stt7/STN7 is a thylakoid associated Ser-Thr regulatory kinase responsible for the phosphorylation of LHCII, which is activated upon reduction of the Cyt b6f (Bellafiore et al., 2005). Stt7 was first identified in mutants of C. reinhardtii with impaired state transitions (Delosme et al., 1996).
Wollman and Lemaire (1988) showed that the Cyt b6f plays a major role in kinase activation in C. reinhardtii. STN7, which is orthologous to Stt7, was first identified in A. thaliana and is conserved in other land plants and in eukaryotic photosynthetic organisms (Bellafiore et al., 2005). Stt7/STN7 kinase has a single transmembrane domain, which separates the small N-terminal region in the thylakoid lumen from the catalytic kinase domain on the stromal side (Lemeille et al., 2009). Apart from being involve in state-transitions, Stt7/STN7 is also involved in long term responses such as adjusting the stoichiometry of the two photosystems to optimize photosynthesis under conditions which favour either one of the two photosystems (Bonardi et al., 2005; Pesaresi et al., 2009). A
90 Stt7/STN7-like protein, Stl1/STN8, is another protein kinase associated with the thylakoid membrane identified in both C. reinhardtii and A. thaliana (Bonardi et al., 2005; Vainonen et al.,
2005; Lemeille et al., 2009). STN8 involved in the quantitative phosphorylation of PSII core proteins (CP43, D1, D2, PsbH), particularly under high light conditions (Bonardi et al., 2005;
Vainonen et al., 2005; Tikkanen et al., 2010). However, studies pointed out that STN8 is not alone in causing PSII core protein phosphorylation and showed a considerable substrate overlap between
STN7 and STN8. This suggested a possible interdependence between these two kinases (Bonardi et al., 2005; Vainonen et al., 2005; Tikkanen et al., 2008; Fristedt et al., 2009). Studies have shown that STN8 also phosphorylates PGRL1 in stn8-1 mutant plants (Reiland et al., 2011) and the chloroplast Ca-sensing protein (CAS) (Vainonen et al., 2008). During dark-light transition, STN8 is thought to cause a rapid switching between CET and linear electron transport (Reiland et al., 2011).
However, the precise role of STN8 in this reversible PSII core phosphorylation has not yet been fully elucidated. A recent study of Wunder et al. (2013) showed that both kinases act in a concentration dependent manner but showed different spatial distributions and modes of regulation.
An early study by Rintamäki et al. (2000) showed that in plants, at high light conditions, the ferredoxin-thioredoxin (Fd-TRX) system inactivates STN7 through the two Cys residues within the
N-terminal domain on the lumen side. It was proposed that CCDA and HCF164 proteins, which are required for haem attachment on the lumen side and for Cyt b6f biosynthesis are also involved in this regulation (Lennartz et al., 2001; Page et al., 2004). However, this effect was not observed in C. reinhardtii at high light (Puthiyaveetil, 2011). A recent study by Wunder et al. (2013) presented experimental evidence for a direct interaction of STN7 and recombinant thioredoxin-f (recΔTRX-f).
This supports the idea of the existence of thioredoxin targeted CxxxC motif in the stromal side of
STN7, which is absent in Stt7, and which is involved in the inactivation of STN7 under high light
(Puthiyaveetil, 2011; Wunder et al., 2013).
91 PPH1/TAP38 is an LHCII specific phosphatase which is involved in the dephosphorylation of
LHCII upon transition from State 2 to State 1 (Pribil et al., 2010; Shapiguzov et al., 2010). This phosphatase dephosphorylates the major trimeric Lhcb1 and Lhcb2 proteins but not the PSII core proteins including, CP43, D1 and D2 (Pribil et al., 2010). PPH1 is a chloroplast protein associated with the stromal membrane and this enzyme belongs to the family of monomeric PP2C type phosphatases (Pribil et al., 2010). Impairment of PPH1/TAP38 results in an increase in the antennae size of PSI and a lack of state-transitions (Pribil et al., 2010). PBPC (PSII core protein phosphatase) is responsible for the dephosphorylation of PSII core proteins, CP43, D1, D2, and PsbH (Samol et al., 2012).
1.5.2. High-energy state Quenching (qE)
High-energy state quenching is the major part of NPQ in plants under high light and relaxes within seconds to minutes (Krause and Weis, 1991; Horton et al., 1994; Horton et al., 1996). qE is defined by a drop of chlorophyll fluorescence quantum yield (Holt et al., 2004). When light absorbed by photosystems exceeds the capacity of electron transport to NADP+ and/or the capacity of ATPase to use the proton gradient to produce ATP, excess protons accumulate in the thylakoid lumen (de
Bianchi et al, 2010). The decrease in pH in the thylakoid lumen or a pH gradient across the thylakoid membrane triggers qE to quench excess light and dissipates it as heat (Horton et al., 1994;
Müller et al., 2001; de Bianchi et al., 2010). A study has shown that application of the ionophore nigericin, disintegrates a pH gradient across the thylakoid membranes thereby, prevents the activation of qE (Ruban and Horton, 1995). Cyclic electron transport (CET) encourages an increase of non-photochemical quenching (NPQ) by increasing the ΔpH across the thylakoid membranes and inhibits the production of reactive oxygen species (Figure 1.9) (Heber and Walker, 1992; Clarke and
92 Johnson, 2001; Golding and Johnson, 2003; Joliot and Johnson, 2011).
Studies on the crystal structure of the isolated LHCII and the antenna proteins have helped to identify the key components important for qE (Sandoná et al., 1998; Liu et al., 2004; Standfuss et al., 2005). They are pigments of the xanthophyll cycle, especially the pigment zeaxanthin (Zea), the
PSII S subunit (PsbS) and the components of the LHCII (Szabó et al., 2005; Wilk et al., 2013).
However, the exact mechanism of qE and the association of these components in the energy dissipation are still under debate (Ruban et al., 2012; Wilk et al., 2013). Several models has been introduced to explain the mechanism of qE. Recent studies have shown that the structural changes occurred within photosynthetic membranes are associated with the mechanism behind qE (Belgio et al., 2014).
The involvement of the xanthophyll cycle on the dissipation of excess energy was proposed more than two decades ago (Frank et al., 1994; Demmig-Adams and Adams, 1996). Violaxanthin (Vio) is a β-carotene-derived xanthophyll pigment, synthesized from Zea through antheraxanthin under low light conditions. This reaction is catalyse by a zeaxanthin epoxidase enzyme (Frank et al., 1994;
Demmig-Adams and Adams, 1996; Szabó et al., 2005). However, under high light conditions, at low lumenal pH, Vio converts into Zea in the presence of Vio de-epoxidase enzyme. A study by
Niyogi et al. (1998) has shown that the npq1 mutants which lack Vio de-epoxidase have low NPQ.
The singlet excited state (S1) of Zea accept energy from excited chlorophyll molecules (Chl*) through singlet-singlet energy transfer and the S1 state of Zea has a short lifetime (10ps) thereby, dissipating energy as heat rapidly (Polivka et al., 2002). The model proposed by Holt et al. (2005) suggested that Zea quenches energy directly and the energy is transferred from excited Chl* to a
Chl-a-Zea heterodimer and forms a Chl-/Zea+ pair through charge separation. Association of PsbS
93 protein with Zea was observed through studies on the optical properties of thylakoid membranes during qE and studies have shown that PsbS is important for direct excitation of Zea and the Chl-a-
Zea heterodimer formation (Aspinall-O'Dea et al., 2002; Ruban et al., 2002; Ma et al., 2003; Holt et al., 2005). Apart from involvement in qE, Zea plays a major role in regulation of the organization of
LHCII-PSII complexes. A study by Havaux et al. (2004) demonstrated that the lut2/npq2,
Arabidopsis double mutant, which is deficient in lutein does not contain any xanthophyll pigments except Zea, which functions as a light-harvesting pigment and is involved in LHCII monomerization, decreasing the size of LHCII and its stability.
The antenna of photosystems consists of specialized membrane- bound light harvesting pigment- protein complexes, where chlorophylls and carotenoids are arranged in a very ordered manner to optimize light capture (Pascal et al., 2005). Both minor LHCII proteins, including Lhcb4 (CP29),
Lhcb5 (CP26) and Lhcb6 (CP24), and peripheral LHCII trimers, including Lhcb1, Lhcb2 and
Lhcb3, are known to be involved in qE (Horton and Ruban, 1992; Wentwort et al., 2004; Horton and Ruban, 2005). However, the mechanisms by which pigments are changed in these complexes to form efficient energy quenchers are yet to identified (Pascal et al., 2005). Binding of dicyclohexylcarbodiimide (DCCD), which inhibits qE by interacting with proton-active residue in the minor LHCII proteins, CP29 and CP26 suggested the involvement of these proteins in qE
(Szabó et al., 2005). Work from Ahn et al. (2008) and Avenson et al. (2008) showed that minor
LHCII subunits are involve in charge transfer quenching. Avenson et al. (2008) used near infra-red
(880-1100nm) transient spectroscopy to show the production of zeaxanthin radical cations in isolated minor LHC complexes, which are involved in charge transfer quenching. Apart from this, in vitro studies showed the occurrence of aggregation and the conformational changes in minor
LHCII during qE (Andersson et al., 2001; Andersson et al., 2003; Dall'Osto et al., 2005).
94 Several models were proposed to show the aggregation of LHCII and proton-induced conformational changes in LHCII associated with pH-dependent quenching. In vitro studies performed by Horton et al. (2005) showed that although quenching does not require oligomerization of proteins, qE is much higher when aggregates formed. However, they also proposed that the formation of LHCII aggregation is not possible in vivo due to the complexity of the thylakoids (Horton et al., 2005). The model suggested by Liu et al. (2004) proposed that, proton-induced conformational change occur in the LHCII trimers, where the site of qE is at the trimer-trimer interface. In this model energy is transferred from chlorophylls to closely located xanthophyll cycle carotenoids (Liu et al., 2004). Another model proposed by Standfuss et al. (2005) showed that the site of qE is located in each LHCII monomer, where chlorophyll-a is close to bound
Vio. Low pH in the thylakoid lumen induces the production of Zea from Vio through xanthophyll cycle. The energy gathered in LHCII monomers transfer energy to the nearby red-shifted chlorophyll-a molecules and then to Zea where energy is dissipated as heat (Standfuss et al., 2005).
However, this model does not suggest the occurrence of any conformational changes in LHCII
(Standfuss et al., 2005; Szabó et al., 2005). Both in vitro and in vivo studies showed that the formation of qE is associated with a change in the configuration of LHCII-bound carotenoid neoxanthin (Robert et al., 2004; Ruban et al., 2007). Changes in the conformation of the components in LHCII change the distance or the orientation between its pigments to form quenching sites (Ruban et al., 2007). Studies by Pascal et al. (2005) and Ruban et al. (2007) pointed out that lutein, the most abundant xanthophyll pigment is responsible for the energy dissipation not
Zea. Betterle et al. (2009) showed that the distances between PSII core complexes decreases under
NPQ. Johnson et al. (2011) and Goral et al. (2012) reported an occurrence of membrane reorganization in intact chloroplasts when exposed to high light which forms clustered domains of
LHCII (monomers and trimers of antenna components and PsbS) and RCII. Miloslavina et al.
(2008) and Holzwarth et al. (2009) showed a functional separation between LHCII and RCII during
95 qE causing a reduction in PSII cross section. They also suggested two NPQ sites: one in the aggregated cluster of LHCII which is detached from the reaction centre core and the other in the light-harvesting complex which is still attached to the core. However, a study of Johnson and Ruban
(2009) and a recent study of Belgio et al. (2014) showed that the functional antenna size of PSII does not decrease during qE.
PsbS (22 KDa) protein was identified as an important component in qE after it was shown that the npq4 Arabidopsis mutants are deficient in qE due to lack of PsbS function (Li et al., 2000).
Although these mutants do not express PsbS protein or express mutated versions, they contain PSII,
LHCII proteins and an active xanthophyll cycle (Li et al., 2000; Peterson and Havir, 2000). A study by Johnson and Ruban (2010) showed that the npq4 Arabidopsis mutant plants lacking PsbS protein possess photoprotective energy dissipation. However, studies have shown that the npq4 Arabidopsis mutants showed a significant reduction in growth under fluctuating light conditions both in the field and in a growth chamber (Külheim et al., 2002; Külheim and Jansson, 2005). Ishida et al. (2011) showed that the suppression of PsbS protein caused a reduction in light-inducible dissipation in rice.
A study by Ikeuchi et al. (2014) showed that PsbS controls the quantum rate of absorbed light energy in PSII allocated to electron transport. Further, they showed that under fluctuating light conditions, the thermal dissipation associated with photoinhibition is enhanced in PsbS-suppressed rice transformants (ΔpsbS). A recent study by Kukuczka et al. (2014) showed that PGRL1 (proton gradient regulation like 1, discussed in section 1.2.2) is a crucial component for PsbS-dependent qE in terrestrial plants to survive under low oxygen and high light conditions. PsbS protein has 8 conserved acidic amino acid (glutamate and aspartate) residues on the lumenal side. These residues are arranged as four symmetrical pairs and are conserved in all photosynthetic species
(Anwaruzzaman et al., 2004). Two of these glutamate residues, E122 and E226 are thought to allow
PsbS to sense low pH levels in lumen which triggers qE (Li et al., 2000; Li et al., 2002; Szabó et al.,
96 2005). Although PsbS was discovered more than three decades ago, its position in PSII and its exact
function in qE has not yet been determined (Teardo et al., 2007; Wilk, 2013). Studies have shown
that this protein is related to Lhcb1-6 proteins (Wedell et al., 1992; Funk et al., 1995). The presence
of PsbS homodimers was identified and it was shown that dimer to monomer transition is triggered
by low pH in the lumen and high light intensities (Bergantino et al., 2003). A recent study by
Haniewicz et al. (2013) pointed out that PsbS protein is bound to PSII monomers in the stromal
lamellae or at the margins of the grana and not the PSII dimers associated with the grana. However,
several studies have suggested that PsbS is strongly associated with the grana (Kiss et al., 2008;
Horton et al., 2008; Kereïche et al., 2010). Several modes of actions of PsbS have been proposed
involving the switching of PSII from a fully active state to a protective state induced by high light
(Haniewicz et al., 2013). PsbS might influence the xanthophyll cycle or might interact directly with
the PSII core or affect the conformational changes in LHCII (Li et al., 2004; Horton et al., 2005;
Szabó et al., 2005; Kiss et al., 2008).
NADPH ATP Fd
CET Stroma
NPQ
S 6 PSII b PQ Cyt b f PSI s P
PC VDE Lumen ATP H+ synthase Low pH Figure 1.9. Regulation of electron transport through qE (reproduced from Shikanai et al., 2014). Under
excess light energy causes the acidification of the lumen through linear electron transport (black
arrows) and cyclic electron transport (dashed black arrows). Low pH is sensed by PsbS and induces qE
component of NPQ and violaxanthin de-epoxidase (VDE), which induces the xanthophyll cycle which
involved in the energy dissipation. Low pH in the lumen reduces the activity of the Cyt b6f complex.
ATP synthase utilises the pH gradients caused by both linear and cyclic electron transport. 97 1.5.3. Other electron transport pathways involved in the regulatory process
+ In addition to the linear electron flow from water to NADP used for CO2 assimilation and photorespiration, there are alternative electron transport pathways in chloroplasts, including the
Mehler reaction, cyclic electron transport around PSI, cyclic electron transport around PSII, chlororespiration and nitrogen assimilation (Heber et al., 1978; Fischer and Klein, 1988; Miyake and Yokota, 2001; Miyake et al., 2002; Makino et al., 2002; Fernandez and Galvan, 2008; Kramer and Evans, 2011). The functions of chloroplasts and mitochondria are closely coordinated, especially under unfavourable environmental conditions (Raghavendra and Padmasree, 2003;
Noctor et al., 2007; Noguchi and Yoshida, 2008). Photorespiration, chlororespiration and N- assimilation are major electron transport pathways associated with both chloroplast and mitochondria. Under stress conditions, reducing equivalents can be transported to mitochondria via the malate and triose phosphate transporters and then used by mitorespiration, thereby increasing
ATP production (Noguchi and Yoshida, 2008). The Pgrl1, a knock out mutant of Chlamydomonas reinhardtii with inhibited cyclic electron flow, showed increased respiration to prevent over- reduction of PSI (Petroutsos et al., 2009). This section aims to discuss the alternative pathways, the
Mehler reaction and chlororespiration. Cyclic electron transport was discussed in Section 1.2.2.
1.5.3.1. Mehler Reaction
The Mehler reaction or the water-water cycle occurs at the acceptor side of PSI and directs excess electrons from PSI to oxygen when PSI acceptors are depleted (Asada, 1999; Asada, 2006).
Photoreduction of molecular oxygen (O2) to hydrogen peroxide in PSI was first reported by Mehler
(1951). It is a pseudo-cyclic electron flow, where electrons from water transferred to PSI and then to
98 oxygen to produce water (Asada, 1999). Electrons accepted by oxygen produce ROS which disproportionates to H2O2 and O2 catalyzed by superoxide dismutase (SOD). H2O2 is further reduced to water by ascorbate peroxidase (APX). Table 1.3 contains the reactions of the water-water cycle
(Asada, 2006).
Table 1.3. Reactions of the Mehler or water-water cycle
Reaction Products Enzymes involved - + 1) 2H2O 4e + O2 + 4H (PSII) - - 2) 2O2 + 2e 2O2 (PSI) - + 3) 2O2 + 2H H2O2 + O2 (PSI) Superoxide dismutase (SOD)
4) H2O2 + 2 ascorbate 2H2O + 2 monodehydroascorbate Ascorbate peroxidase (APX) (AsA) radical (MDA) (PSI) 5) 2MDA + 2 reduced Fd 2 AsA + 2Fd (PSI) Occurred spontaneously 6) 2MDA + NAD(P)H 2AsA + NAD(P)+ (PSI) MDA reductase 7) 2MDA AsA + dehydroascorbate Disproportionates into AsA and DHA (DHA) (PSI) 8) DHA + 2 reduced AsA + 2GSH (PSI) DHA reductase and glutathione glutathione (GSH) reductase 9) 2 Fd or NAD(P)+ + 2e- reduced Fd or NAD(P)H (PSI)
Apart from a possible photoprotective role, the Mehler reaction has also been suggested to be important to generate a pH gradient across the thylakoid membranes when PSI acceptors are depleted which is necessary for NPQ (Osmond and Grace, 1995; Osmond et al., 1997; Asada, 1999;
Asada, 2000; Foyer and Noctor, 2000). However, a study by Makino et al. (2002) showed that although the Mehler reaction is important to build a pH gradient across the thylakoid membranes which initiates NPQ and ATP production, CET is the major pathway which is responsible to maintain high NPQ. Although the Mehler reaction has been argued to act as a safety process, to divert excess electrons from the electron transport, it has several drawbacks which outweigh any potential benefits. Reduction of molecular oxygen forms harmful radical species which can damage
99 PSI and PSII (Asada, 1999; Asada, 2000; Asada, 2006). Apart from that, producing scavenging enzymes in higher concentrations is energetically demanding for plants. According to several studies, the Mehler reaction is insufficient to provide significant protection from photoinhibition
(Cornic and Briantais, 1991; Wiese et al., 1998; Clarke and Johnson, 2001; Driever and Baker,
2011).
1.5.3.2. Chlororespiration
Evidence has been found regarding the existence of an alternative electron transport pathway which is a thylakoid associated respiratory chain which has components homologous to those involved in mitochondrial respiration and this is called chlororespiration (Nixon, 2000; Peltier and Cournac,
2002). Chlororespiration is described as a light independent electron transport pathway in the chloroplast involving plastoquinone as an electron carrier (Houille-Vernes et al., 2011). In the early
1960's, a study from Goedheer (1963) suggested dark oxidation of intersystem electron carriers occurred via “some kind of chloroplast respiration” by analysing luminescence transients in unicellular green algae. In 1982, Bennoun, based on the effects of respiratory inhibitors on chlorophyll fluorescence induction curves in unicellular green algae, proposed the existence of respiratory chain associated with the thylakoid membrane, which is connected to the photosynthetic electron transport chain (Bennoun, 1982).
Although the occurrence of chlororespiration was under debate in the past, after the discovery of two membrane-bound proteins, plastid-encoded NAD(P)H-dehydrogenase (NDH) and nuclear- encoded plastid terminal oxidase (PTOX), the potential for chlororespiration now appears to be well established in both algae and higher plants (Peltier and Cournac, 2002). According to the current model of chlororespiration, reduction of PQ occurs through NAD(P)H-dehydrogenase and plastid
100 terminal oxidase diverts electrons from the PQ pool to molecular oxygen (Nixon, 2000; Peltier and
Cournac, 2002). Both of these proteins have been shown to be located in the nonappressed membrane (Berger et al., 1993; Sazanov et al., 1996; Horváth et al., 2000; Lennon et al., 2003;
Kuntz et al., 2004). Therefore, chlororespiration is proposed to be restricted to the stromal lamellae of the chloroplast (Peltier and Cournac, 2002). Early studies showed that type I chloroplastic NDH complex is absent in many green alga including Chlamydomonas (Peltier and Cournac, 2002;
Robbens et al., 2007). However, Jans et al. (2008) showed that Chlamydomonas thylakoids contain a type II NAD(P)H dehydrogenase (NDH-2 or NDA2) which mediates the light-independent PQ reduction in the thylakoid membrane. The NDH complex which is involved in both CET and chlororespiration consists of a large number of chloroplast- and nuclear-encoded subunits (Rumeau et al., 2007). A study from Houille-Vernes et al. (2011) showed that plastid terminal oxidase 2
(PTOX2) is the major plastoquinol oxidase involved in chlororespiration.
Early experiments were based on assumptions that the chloroplast and mitochondria were not in redox communication with each other (Bennoun, 1982). However, both mitochondria and chloroplast possess a metabolic interaction through exchanging reducing equivalents like
NAD(P)H, or the phosphorylating power of ATP (Raghavendra et al., 1994; Gardeström and
Lernmark, 1995; Hoefnagel et al., 1998). This is performed by metabolic shuttles, such as the phosphate/dicarboxylate translocators in the inner chloroplast envelope and the oxaloacetate translocators in the inner mitochondrial membrane (Heber, 1978; Heldt et al., 1990; Raghavendra et al., 1994; Hoefnagel et al., 1998). The interaction between mitochondria and chloroplast is observed in Chlamydomonas chloroplast ATPase mutant which is unable to produce chloroplast ATP and rely on ATP produced in the mitochondria (Lemaire et al., 1988). Another study has shown that inhibition of mitorespiration decreases cellular ATP production, encouraging glycolysis in the chloroplast resulting in rising levels of NAD(P)H and rates of reduction of the PQ pool through
101 NAD(P)H dehydrogenase (Gans and Rebeillé, 1990). Similar to photosynthetic purple bacteria, it is proposed that mitochondrial oxidases oxidize the PQ pool through reverse electron flow from PQ to
NAD(P)+ which is also catalysed by NAD(P)H dehydrogenase and driven through an electrochemical gradient across membranes (Bennoun, 1994).
Due to the interaction between chloroplast and mitochondria it was difficult to differentiate the activity of mitochondrial oxidases and chloroplast oxidases involved in the PQ oxidation (Bennoun,
1994; Bennoun, 1998; Bennoun, 2001). However, studies from Cournac et al. (2000) and Bennoun
(2001) showed the presence of plastoquinol: oxygen oxidoreductase in the chloroplast membranes responsible for dark oxidation of the plastoquinol pool. Cournac et al. (2000) discovered a PSII- dependent O2 production and O2 uptake in isolated chloroplast fractions of Chlamydomonas mutant
18 deficient in PSI by using continuous mass spectrometry in the presences of O2. This is insensitive to many inhibitors including azide, carbon monoxide, cyanide, antimycin A and salicylhydroxamic acid (SHAM) but sensitive to propyl gallate (PG) (Cournac et al., 2002). Work from Bennoun
(2001) showed that plastoquinol oxidation was inhibited in Chlamydomonas mutants deficient in the cytochrome b6f complex when the algal cells were exposed to low O2 concentrations (mixture of air and argon at 1.45% air). The study also showed that plastoquinol oxidation in darkness is sensitive to n-propyl gallate (PG) but not to SHAM, whereas mitochondrial respiration is sensitive to both PG and SHAM) (Bennoun, 2001). These experimental evidences led to the conclusion of the involvement of a chloroplast oxidase which is responsible for the oxidation of PQ pool. This is now known as plastid terminal oxidase (PTOX) (Cournac et al., 2000; Cournac et al., 2002).
102 1.6. Involvement of Plastid terminal oxidase (PTOX) in alternative electron transport in the electron transport chain of Thellungiella salsuginea
Persistent stress acts as a selective factor and triggers plants to establish resistance and improve adaptability. The recovery processes during stress conditions causes hardening of plants to cope with the ever changing climate. Beck et al. (2004) found that low temperature and short photoperiods enhance frost hardening in Scots pines (Pinus sylvestris L) during autumn. However, most plants suffer from challenging weather and poor soil conditions worldwide. Abiotic stresses are one of the major reasons for crop losses every year. For example rice is considered to be salt- sensitive and production decreases every year, due to increasing salinity of the soil (Khatun and
Flowers, 1995). Therefore, producing stress tolerant crop varieties is the only possible solution to improve the crop production in the marginal lands. Some wild plants which grow in poor soil and adverse environmental conditions, known as extremophiles, contain important genes which are involved in stress tolerance and might be transferred in to crop varieties to enhance the vigor and improved productivity (Amtmann, 2009). Plants, including Thellungiella salsuginea and Xerophyta viscosa are such extremophiles, studied because of their highly tolerant nature to abiotic stresses
(Volkov and Amtmann, 2006; Kant et al., 2008; Lehner et al., 2008; Amtmann, 2009; Wu et al.,
2012).
Thellungiella salsuginea (common names: salt-water cress and salt-lick mustard) is a halophyte, which is tolerant to many abiotic stresses including salinity, cold, drought and nitrogen poor soils
(Griffith et al., 2007; Amtmann, 2009). It belongs to the family Brassicaceae and is closely related to Arabidopsis thaliana (sequence identity of 92%). T. salsuginea has short life cycle and small genome, which has recently been sequenced (Amtmann, 2009; Orsini et al., 2010). A short life cycle, copious seed production and an ability to grow in laboratory conditions have made T.
103 salsuginea one of the valuable model plants for research on abiotic stress tolerance (Bressan et al.,
2001; Inan, 2004; Wong et al., 2005; Wong et al., 2006).
T. halophila, T. salsuginea and T. parvula are the identified species of Thellungiella (Amtmann,
2009; Pedras and Zheng, 2010). However, T. salsuginea is also known Arabidopsis halophila,
Arabidopsis glauca, Eutrema halophilum, Eutrema salsugineum and Arabidopsis salsuginea (Al-
Shehbaz et al., 1999; Griffith et al., 2007; Stepien and Johnson, 2009). T. salsuginea has two studied ecotypes: Shandong (native to China), which was the first identified and studied and Yukon (native to Canada). Other ecotypes of Thellungiella species are distributed in various habitats with extreme environmental conditions, from the subarctic, arctic, alpine regions to saline meadows (Griffith et al., 2007; Amtmann, 2009).
Although T. salsuginea is morphologically similar to A. thaliana, the leaves have serrated margins
(Amtmann, 2009). T. salsuginea leaves have densely distributed epicuticular wax crystals compared to the leaves of A. thaliana (Teusink et al., 2002). A study by Inan et al. (2004) showed that T. salsuginea has high stomatal density distributed over the surface but not fully open compared to A. thaliana. Leaves have a second layer of palisade mesophyll cells which is shed during extreme salt stress and roots have thicker endodermis and cortex cell layers compared to A. thaliana (Inan,
2004). The leaf surface of Thellungiella is glaucous throughout development, whereas, in
Arabidopsis, leaf surface is more hairy (Teusink et al., 2002).
T. salsuginea can grow and reproduce in extreme salt concentrations (500mM) and freezing temperatures (-19 oC) (Inan, 2004; Gong et al., 2005; Griffith et al., 2007; Stepien and Johnson,
2009). Work from Griffith and co-workers (2007) pointed out that T. salsuginea lacks endogenous ice nucleation and therefore managed to survive in freezing temperatures. Wang and co-workers
104 (2013) showed that T. salsuginea is able to survive in extreme saline soils by compartmentalizing
Na+ into the cell vacuoles. In addition to that, T. salsuginea plants accumulate proline and soluble sugars which are used as osmolytes under water deficit conditions. Unlike A. thaliana, T. salsuginea can lives in nitrogen-limiting conditions. T. salsuginea grown in nitrogen-limiting conditions showed low carbon to nitrogen ratio, high nitrogen content, high total amino acid content, high total soluble proteins, low starch content, high soluble sugars and high organic acids content compared to
A. thaliana (Kant et al., 2008). It was found that T. salsuginea produces various antimicrobial substances including phytoalexins and phytoanticipins when exposed to abiotic and biotic stresses
(Pedras and Zheng, 2010).
Stepien and Johnson (2009) found that, T. salsuginea showed an increase in the PSII electron transport rate when plants were exposed to 250 mM salt concentrations, whilst this was not seen in the salt-treated A. thaliana. However, this effect was not observed when the oxygen level was decreased to 2%. The study also showed the PSI ETR was unaltered, suggesting the oxygen sensitive electron transport does not involve PSI. According to immunoblot analysis it was found that a protein reacting against antibodies to plastid terminal oxidase (PTOX) is more prominent in
T. salsuginea than A. thaliana. From this study they concluded that, in salt-stressed T. salsuginea, plastid terminal oxidase acts as an alternative electron sink, accounting for up to 30% of total PSII electron flow.
Plastid terminal oxidase (PTOX, also known as plastoquinol terminal oxidase) is a chloroplast targeted terminal plastoquinol oxidase, which plays a vital role acting as an alternative electron sink and directing excess electrons to oxygen to produce water. PTOX is produced in plastids and is homologous to mitochondrial alternative oxidase (AOX). Based on gene sequence analysis, the predicted structure of the PTOX protein has a di-iron carboxylated centre in the active site with 6-
105 ligands (4 glutamates and 2 histidines), which is similar to that of AOX (Berthold and Stenmark,
2003; McDonald et al., 2011). PTOX is proposed to be an interfacial membrane protein which is encoded by a single nuclear gene in higher plants and two genes (PTOX1 and PTOX2) in prasinophytes, chlorophytes, diatoms and red alga and sometimes significantly divergent, PTOX paralogues (Wang et al., 2009). A study by Houille-Vernes (2011) showed that in C. reinhardtii,
PTOX2 is the major oxidase involved in chlororespiration.
PTOX was first identified in an A. thaliana pigment mutant known as immutans and later identified in the tomato ghost mutant (Wu et al., 1999; Josse et al., 2000; Carol and Kuntz, 2001). It is known to be involved in phytoene desaturation in the carotenoid biosynthesis pathway. PTOX mutants, immutans and ghost produce a variegated phenotype under low-medium light and, under strong light, leaves bleach, due to oxidative stress (Carol et al., 1999; Wu et al., 1999; Kuntz, 2004).
Variegated plants consist of green and white/yellow sectors in most of the green organs of the plant.
Compared to the green sectors, cells in white sectors have chloroplasts which lack pigments. These studies revealed that PTOX plays a major role in lowering the excitation pressure in PSII during carotenoid biosynthesis at the early stages of chloroplast biogenesis (Carol et al., 1999; Aluru et al.,
2006; Rosso et al., 2009; Foudree et al., 2012). Okegawa et al. (2010) showed that impairment of
PSI cyclic electron transport suppressed leaf variegation in the A. thaliana immutans, which is deficient in PTOX. Inhibition of carotenoid biosynthesis caused interruptions in other processes such as synthesis of ABA (Aluru et al., 2001). A recent study on the ptox1 mutant of Oryza sativa showed that PTOX1 is required for both carotenoid and strigolactones synthesis (a family of plant hormones that are synthesized from carotenoids) (Tamiru et al., 2014).
Involvement of PTOX in chlororespiration and the regulation of the redox state of the PQ pool have been extensively studies over last few years (Peltier and Cournac, 2002; Aluru and Rodermel,
106 2004). PTOX involved in the oxidation of the PQ pool under dark-to-light transition when the
Calvin cycle is not yet activated. Plants including Rananculus glacialis, T. salsuginea, Brassica fruticulosa and Brassica oleracea showed an increase in both PTOX activity and protein levels under abiotic stresses (Streb et al., 2005; Díaz et al., 2007; Stepien and Johnson, 2009; Laureau et al., 2013). Apart from this increase in PTOX mRNA transcripts were also observed in plants like
Coffee arabica and Oryza sativa under drought and salinity, respectively (Kong et al., 2003; Simkin et al., 2008). Shirao et al. (2013) found that, gymnosperms have increased capacity for electron leakage to oxygen (Mehler and PTOX reactions) in photosynthesis compared with angiosperms.
However, overexpression studies on A. thaliana and tobacco has shown that PTOX did not confer enhanced protection against photoinhibition in these plants (Joët et al., 2002; Rosso et al., 2006;
Ahmad et al., 2012). A recent study by Yu et al. (2014) showed that PTOX protein exists mainly as a homo-tetrameric complex with two Fe per monomer and is very specific for a plastoquinone head group. Further, the study concluded that PTOX can act as a safety valve when the steady state PQH2 is low while a certain amount of ROS is formed at high light intensities.
Apart from plants, activity of PTOX under various stress conditions were observed in other photosynthetic organisms. It was found that a PTOX gene is present in all high-light adapted ecotypes of Prochlorococcus marinus but not in the cyanobacterial strains found in low light and low temperature environments (Rocap et al., 2003; Kettler et al., 2007; Luo et al., 2008). A study showed that Synechococcus WH8102, a marine cyanobacterium possess an alternative electron flow to O2 via PTOX when PSI activity is limited due to low iron levels (Bailey et al., 2008). They hypothesized that Synechococcus uses PTOX, which only has two iron atoms rather cytochrome b6f and PSI which have 18 iron atoms altogether, to survive in low iron conditions. Apart from that, expressed sequence tag data suggested that PTOX was transcribed in two diatoms Phaeodactylum tricornutum and Thalassiosira pseudonana under low iron conditions (McDonald et al., 2011).
107 Studies on green algae, Haematococcus pluvialis showed that changes in the PTOX transcripts under various stresses including high light, excess iron or salt and low temperature (Li et al., 2008;
Wang et al., 2009; Li et al., 2010). They also showed that PTOX involved in the production of astaxanthin and plays a protective role against stress (Li et al., 2008; Wang et al., 2009; Li et al.,
2010). Work from Cardol et al. (2008) showed a deep sea/low light strain of the green, picoeukaryote Ostreococcus strain (RCC 809) lives in low iron conditions lacks PSI compared to surface/ high light strain (OTH95) bypasses electrons in a water-water cycle to generate a pH gradient across the thylakoid membranes. This cycle bypasses large number of electrons generated through PSII to oxygen with the involvement of PTOX (Cardol et al., 2008). Increased levels of
PTOX transcripts in phosphorus starved cells of C. reinhardtii suggested that PTOX plays a major role in stress responses in this photosynthetic algae (Moseley et al., 2006). Apart from the photosynthetic organisms, cyanopages, viruses which infects cyanobacteria, possess PTOX genes in their genomes. For example, cyanophage Syn9 consists of photosynthetic genes including, plastocyanin, PTOX, PsbA, PsbD (Weigele et al., 2007). It has been hypothesized that these genes provided a photoprotective function during the phage propagation (Lindell et al., 2005; Weigele et al., 2007).
Although researchers have proposed several possible pathways of PTOX mediating alternative electron transport, the definite pathway is yet to be identified. The one possible pathway is a direct transport of electrons from the PQ pool through PTOX to oxygen to produce water. Other suggested pathway involves the NDH complex (NAD(P)H dehydrogenase) known to be involved in cyclic electron transport. Electron transfer might pass through the NDH complex and then to oxygen via
PTOX to produce water (Kuntz, 2004; McDonald et al., 2011). It is evident that in some plants,
PTOX act as an alternative sink or a safety valve to remove excess electron gathered around PQ pool (Streb et al., 2005; Díaz et al., 2007; Stepien and Johnson, 2009; Laureau et al., 2013). This
108 prevents the formation of ROS and protects the photosynthetic apparatus. However, this protective mechanism is not common in the whole plant kingdom (Sun and Wen, 2011). Therefore, coupling the level and the activity of PTOX with other photo-protective mechanisms in stress sensitive plants may help them to survive in an ever changing environment.
109 1.7. Aims and Objectives
The aim of this project is to understand the physiology of photosynthesis in stress tolerant and stress sensitive plants and to characterize the tolerant traits in plants which are responsible for the regulation of photosynthesis under abiotic stress. Plants such as Thellugiella salsuginea are tolerant to abiotic stresses, whereas other plants, including most crops, are sensitive to changing environmental conditions. Stress tolerant plants incorporate traits which are important to regulate photosynthesis under stress. Therefore, it is important to understand the physiology of stress tolerance and characterise stress tolerance traits in plants.
The first experimental chapter focuses on analysing the effects of salt stress on both photosystems in barley. Both PSII and PSI electron transport was measured, along with gas exchange and chlorophyll content. The aim of this chapter is understand the effects of salt stress on the electron transport of barley variety Chalice at the early vegetative stage.
The second data chapter provides a physiological evaluation of salinity stress in two rice varieties from Sri Lanka. The aim of this chapter is to compare the physiology of photosynthesis in salt- tolerant and salt-sensitive rice and to characterize the salt-tolerant traits which are responsible for the regulation of photosynthesis. Sri Lanka, as one of the major rice producing countries of the world, loses most of its production due to salinity every year. Researchers are mainly focusing on characterizing traits which help plants to regulate metabolic processes in order to survive in saline environmental conditions and produce salt-tolerant rice varieties. PSII, PSI, gas exchange, leaf area and chlorophyll content of both early vegetative and flowering stages were measured under salt stress. Differences in the regulation of the electron transport chain are also discussed in this chapter.
110 The third experimental chapter is focused on the activity of the plastid terminal oxidase (PTOX) and the regulation of T. salsuginea under abiotic stresses. According to Stepien and Johnson (2009),
PTOX activity is observed in T. salsuginea under salt stress. Therefore, in this chapter measurements were carried out to examine the presence of PTOX under other abiotic stresses including drought, low temperatures and different growth irradiances. Further analyses were done to examine the transcriptional regulation of PTOX gene. This was performed by detecting the mRNA transcripts in stressed plants. In addition to that, further analyses were performed to determine the possible location of PTOX protein on the thylakoid membrane and possible associated complexes.
111 Chapter 2
Effects of salt stress on the regulation of photosynthesis in
barley
(Hordeum vulgare L.)
Sashila Abeykoon Walawwe
Giles N. Johnson
112 Preface
Sashila Abeykoon Walawwe is the primary author of this paper.
Plant growth by Sashila Abeykoon Walawwe
Measurements of chlorophyll content by Sashila Abeykoon Walawwe
Photosynthetic measurements of PSII and PSI by Sashila Abeykoon Walawwe
113 2.1. Abstract
Salinity is one of the important environmental stresses which adversely affects the productivity of crops worldwide. Barley is considered as a crop which is comparatively tolerant to soil salinity.
Much research had been done to characterise the salinity tolerance in barley. The focus of this study is to evaluate the physiological responses of photosynthesis in barley under salinity.
Barley cultivar Chalice was grown hydroponically and treated with 50, 100 or 250 mM NaCl concentrations. 14 days after the start of salt treatment, the effects of salt stress on gas exchange, chlorophyll fluorescence, PSI electron transport and chlorophyll content were examined. Two-week exposure to salt decreased the rate of CO2 assimilation, stomatal conductance, the rate of transpiration, electron transport rate of PSII (PSII ETR) and electron transport rate of PSI (PSI
ETR) and chlorophyll content of barley. At low salt concentrations, barley plants protect themselves by down-regulating the photosynthetic electron transport chain. At 250 mM, barley showed a significant decrease in PSII ETR and the rate of CO2 assimilation. Although PSII ETR decreased with high salt concentrations, PSI ETR was not substantially reduced with increasing salt concentration. From this, it seems likely that cyclic electron transport is enhanced in salt-stressed barley. Cyclic electron transport helps to maintain a pH gradient required to support non- photochemical quenching (NPQ). However, at 250 mM NaCl cyclic electron transport fails to regulate electron transport. This might be due to loss of PSI centres or an increase in leakiness in the thylakoid membranes.
114 2.2. Introduction
Excess salt in soil and water affect plants by disturbing the osmotic potential in cells and causing toxicity and/or nutritional disorders (Läuchli and Epstein, 1990; Läuchli and Grattan, 2007).
Therefore salinity causes perturbations in all physiological and biochemical processes in plants
(Parida and Das, 2005). Photosynthesis is one of the major metabolic processes severely affected by salinity and plants show complex photosynthetic responses to salt stress (Chaves et al., 2009). Soil salt directly impacts on photosynthetic pigments, proteins in the thylakoid membrane, electron transport reactions, photophosphorylation and CO2 fixation (Kalaji and Nalborczyk, 1991; Delfine et al., 1999; Sudhir et al., 2005). Soluble proteins in leaves also decrease when plants are exposed to high salt concentrations (Alamgir and Ali, 1999; Gadallah, 1999; Wang and Nii, 2000; Parida and
Das, 2005). At high salt concentrations, plant roots are unable to absorb water from the soil. Plants prevent water loss through transpiration by closing stomata. As a result, the entry of CO2 into the leaf is restricted which inhibits CO2 assimilation (MacRobbie, 1998; Blatt, 2000; Hetherington and
Woodward, 2003; Shimazaki et al., 2007; Kim et al., 2010). Studies have shown that mesophyll conductance is an important factor under salinity because it affects the CO2 diffusion into cells
(Centritto et al., 2003; García-Sánchez and Syvertsen, 2006). Most higher plants, such as tomato, potato, pea and bean, are highly susceptible to salt stress and show decreases in chlorophyll content
(Lapina, 1970; Seemann and Critchley, 1985; Abdullah and Ahmad, 1990).
Salt induced responses and tolerance mechanisms in plant depends upon many factors, including the species, genotype, plant age, ionic strength and composition of the salinizing solution, and the organ in question (Läuchli and Grattan, 2007). Salinity tolerance in dicotyledonous species is more prominent than amongst monocotyledonous plants (Munns and Tester, 2008). Barley (Hordeum vulgare L.) is the fourth most widely grown crop and is used as an animal fodder and as raw
115 material for alcohol production (Schulte et al., 2009; Widodo et al., 2009). Compared to other cereals, such as rice and wheat, barley is a relatively salt tolerant crop (Hayes et al., 1996; Colmer et al., 2005; Munns and Tester, 2008; Witzel et al., 2009, Gupta and Huang, 2014). Barley is considered as a salt includer (translocates Na+ to the shoot rather than retaining it in the roots) and is sometimes described as a halophyte (Glenn et al., 1999). The salt tolerant nature of barley makes it an important model plant to examine salt tolerance traits, which might be incorporated into other salt sensitive cereals (Mian et al., 2011). However, some studies have shown that there is a wide variability among barley cultivars (Epstein and Norlyn, 1977; Rathore et al., 1977; Day et al., 1986;
Morales et al., 1992; Forster et al., 2000) and species (Mano and Takeda, 1998) in resistance to salinity. Belkhodja et al. (1999) showed that cultivar Albacete is less affected by high salt concentrations than the other cultivars, such as Dacil and Igri. Widodo et al. (2009) showed that a barley variety known as Sahara has relatively high leaf Na+, less necrosis and high salt tolerance compared to the variety Clipper. Janušauskaitė et al. (2013) showed contrasts in gas exchange parameters, including the rates of assimilation and transpiration, stomatal conductance and instantaneous water use efficiency between different barley varieties.
Salt tolerance in barley is influenced by genetic diversity and the adaptation to a broad spectrum of micro-ecological conditions (Nguyen et al., 2013). Barley has a rich genepool with a large variation in adaptation to abiotic stresses such as drought and salinity (Nevo and Chen 2010). Because of that, barley is considered as a source of favourable alleles to be used in cereal salt tolerance improvement through conventional and molecular approaches (Colmer et al., 2006; Munns et al.,
2006). Physiological, genetic and cytogenetic studies were performed to understand the salt tolerance of barley (Cramer et al., 1991; Forster et al., 1997; Mano and Takeda, 1997; Munns and
Rawson, 1999; Munns et al., 2000; Ellis et al., 2002; Tavakkoli et al., 2010). Transcriptional profiling on barley cultivar, Morex was performed to analyse the early responses of genes to salinity
116 stress at the seedling stage (Walia et al., 2006). This study found that many genes involved in jasmonic acid biosynthesis pathway were induced under salt stress, suggesting the involvement of osmoprotectants at the early stage of barley against salinity. A recent study on quantitative trait loci
(QTL) of barley (collection of 192 genotypes from a wide geographical range) showed a large variation of traits that were highly heritable under salt stress and these traits contribute to salt tolerance (Nguyen et al., 2013). A study on the salt-induced root proteome of barley emphasized the expression of proteins involved in ROS detoxification during salinity stress in the tolerant genotype.
However, in the sensitive genotype, proteins which are involved in ion uptake were expressed abundantly (Witzel, 2009).
Although barley is considered as a relatively resistant crop variety to salt, Royo and Aragüé (1993) and Sánchez-Díaz et al. (2002) found that barley grown under salt stress showed a marked reduction in shoot and root biomass, resulting in total yield loss. Work from Belkhodja et al. (1999) showed that, soil salt reduces the leaf size, but increases the stomatal frequency in barley. In this study, the net photosynthetic rate and the stomatal conductance to water vapour in flag leaves of barely showed a distinct reduction at high salt concentrations. The distinct correlation of stomatal
conductance may suggests that the reduction of CO2 assimilation rate is due to the closure of stomata. Therefore, the closure of stomata due to the high salt concentrations is the most important effect for the retardation of photosynthesis in barley (Belkhodja et al., 1999). Kalaji et al. (2011) found that, the first stage of salinity effects on photosynthesis of barley are related to stomatal limitation rather than to PSII activity reduction. Reduction in the efficiency of PSII in salt treated barley is suggested to be due to D1 protein degradation and the inactivation of PSII reaction centres
(Kalaji et al., 2011). A study by Pérez-López et al. (2012) showed that elevated CO2 levels reduces stomatal and non-stomatal limitations on photosynthesis of salt-treated plants of the barley cultivar
Iranis. In some plants, salinity changes maximum quantum efficiency, Fv/Fm measured using
117 chlorophyll fluorescence. However, the rapid kinetics of chlorophyll fluorescence have indicated that the PSII photochemical efficiency of barley was not affected by high salt concentrations even when the growth is highly retarded (Morales et al., 1992). This effect is also seen in barley when exposed to drought and it was shown that cyclic electron transport occurred to reduce the excitation pressure built up in drought stressed plants (Golding and Johnson, 2003).
Although, the negative effects of salt on photosynthesis have been known for a long time, these are not yet fully understood (Kalaji and Łoboda, 2009; Kalaji et al., 2011). There is a wide variability among barley varieties in resistance to salt and it is not yet clear whether there is one common salt tolerance mechanism in different cultivars (Kalaji et al., 2011). A number of studies provided important information on the effects of salt stress on the electron transport chain of photosynthesis in barley through simultaneous measurements of gas exchange and chlorophyll fluorescence (Jiang et al., 2006; Tavakkoli et al., 2010; Kalaji et al., 2011; Pérez-López et al., 2012; Kalaji et al., 2013).
However, it will be useful to measure the effects of salt of PSI electron transport along with gas exchange and chlorophyll fluorescence measurements, to identify regulatory processes occurring in the electron transport chain in response to salinity. Therefore, the goal of the current study was to assess effects of different salt concentrations on gas exchange parameters, chlorophyll fluorescence,
PSI photochemistry and chlorophyll content in barley cultivar Chalice.
118 2.3. Methods and Materials
2.3.1. Plant growth
Seeds of barley (Hordeum vulgare L.) variety Chalice were germinated on water-saturated tissue paper in a sealed translucent container at 22 oC in a growth cabinet at a light intensity of 140
µmolm-2 s-1 provided from white fluorescent bulbs. After 5 days, seedlings were transferred to a hydroponics system, containing a nutrient medium (NitrozymeTM, 5 ml per 10 litres of water). The nutrient solution was changed 3 times a week and plants were grown in this solution for 7 days.
Then plants were transferred to a nutrient solution containing 0, 50, 100 or 250 mM NaCl for up to
14 days.
The first leaf from 4 barley plants at the early vegetative stage were used for measurements of gas exchange, chlorophyll fluorescence, PSI electron transport and chlorophyll content. Measurements were repeated four times for each salt treatment and measurements being performed on the leaf between 4 and 7 cm from the leaf tip.
2.3.2. Measuring Photosynthetic parameters (Gas exchange, P700 oxidation and chlorophyll fluorescence measurements)
2.3.2.1. Measuring gas exchange
The measurements of gas exchange were performed in combination with chlorophyll fluorescence analysis. Gas exchange measurements were taken using an infra-red gas analyser (IRGA; CIRAS-1,
119 PP systems Ltd, Herts, UK). Leaves were clamped side by side in the cuvette chamber to fill the chamber without overlapping. Temperature and humidity were maintained at 25 oC and 60%, respectively. Leaves were left to equilibrate in the dark for 5 minutes. The rate of gas exchange was measured for 1-2 minutes. After that leaves were illuminated for 20 minutes until a stable rate of gas exchange was obtained (Johnson and Murchie, 2011). The external CO2 concentration was maintained at 390 µL L-1 and assimilation rate (A), stomatal conductance to water vapour (gs) and transpiration rate (T) were measured for each actinic light intensity. Actinic light (up to 1, 600 µmol m-2 s-1) was supplied by a Luxeon III red LED in a laboratory built lamp and light intensity was measured using a PAR meter (SKP215; Skye Instruments, Powys, UK).
A gas with known concentration of CO2 and water vapour was passed through the chamber at a constant 200 cm3/minute rate. Differences in absorbance in the infra-red (IR) analyser were used to
measure the amount of CO2 and water in gas leaving the chamber, compared to that entering. The flux in CO2 and water per unit leaf area were calculated by measuring the differences in gas concentration between the reference line and the leaf sample in the cuvette chamber. The humidity in the internal leaf air spaces, the leaf temperature and the external leaf humidity were used to calculate the total conductance of the leaf to water vapour. This is calculated using the Fick's law of diffusion (Caemmerer and Farquhar, 1981). The stomatal conductance to water vapour was calculated by removing the boundary layer contribution (as estimated by the manufacturer for a given chamber). The CO2 conductance is obtained from the stomatal conductance by correcting for the physical properties of CO2 and water molecules. Internal CO2 concentration, Ci, is the sub- stomatal or mesophyll cell wall CO2 concentration. Ci, is assumed to be the in vivo substrate concentration of Rubisco, calculated using the CO2 conductance, assimilation rate and transpiration rate (Caemmerer and Farquhar, 1981; Johnson and Murchie, 2011).
120 An ACi curve can be used to separate the various limiting steps in photosynthesis, including
Rubisco activity, RuBP regeneration, triose phosphate utilization and stomatal limitations (Farquhar and Sharkey, 1982; Johnson and Murchie, 2011). Measurements for ACi curves were taken under saturating actinic light intensity of 1000 µmol m-2 s-1. First, leaves in the cuvette chamber was
-1 equilibrated with ambient CO2 level (390 µL L CO2). Then leaves were exposed to series of CO2 concentrations, starting from low values (typically minimum of between 10 and 40 µL L-1). The time lag between two measurements at different CO2 concentrations was restricted to 4 minutes and each curve was completed within 20-30 minutes (Johnson and Murchie, 2011; Pérez-López et al.,
2012). CIRAS-1 software was programmed to calculated Ci value for each CO2 concentrations used.
2.3.2.2. Chlorophyll Fluorescence and PSI measurements
Changes in absorbance at 830-870 nm were used to give a measure of the redox state of the PSI primary donor, P700 (Harbinson and Woodward, 1987; Klughammer and Schreiber, 1994).
Measurements were made using a Walz PAM 101 fluorometer in combination with an ED-
P700DW-E emitter-detector unit (Walz, Effeltrich, Germany). Actinic light (up to 1, 600 µmol m-2 s-
1) was supplied by a Luxeon III red LED in a laboratory built lamp. The data were observed and captured using a National Instruments PCI-6220 data acquisition card, in a computer running software written using Labview (National Instruments, Austin, TX, USA). Chlorophyll fluorescence measurements of PSII were made using a PAM 101 fluorometer together with a 101-ED emitter- detector unit (Walz). Fluorescence data were recorded and captured using the same software
(Golding and Johnson, 2003; Hald et al., 2008).
For measurements on any given leaf sample, the following sequence of procedures were carried out:
121 1. Maximum P700 signal
To measure the signal corresponding to 100% of P700, dark-adapted leaves were exposed to far-red light, maximum 730 nm provided by a far red LED array (Roithner Lasertechnik, Vienna, Austria) for 1 minute. To check whether the light was saturating, a 100 milliseconds flash of red light (4000
µmol m-2 s-1) was applied on top of the far-red light. The saturation of the far-red light was indicated by the absence of the substantial rise of the signal after applying the flash (Golding and Johnson,
2003). This gives the maximum oxidation of P700 (P700total) (Figure 2.1).
SP on Signal size for P700total
FR on
Figure 2.1. Far-red light (FR) induced signal giving the 100% of P700. Leaves were exposed to far- red light, λmax = 730 nm, for 1 minute. A 100 milliseconds flash of red light (SP) (4000 µmol m-2 s-
1) was applied on top of the far-red light to check whether it is saturated. The absence of the a rise in the signal after applying the red flash indicates that the signal was saturated.
122 2. Chlorophyll fluorescence
Leaves were allowed to recover for two minutes after illuminating with FR light (to re-reduce all the P700+). Fo (zero level of fluorescence) was measured by applying the PAM-101 measuring light for 10-20 seconds. A high intensity short duration saturating light (1 second pulse of red light with a
PFD of 4000 µmol m-2 s-1) was applied to reached Fm (maximum fluorescence). Following that, actinic light was switched on and plants were left for 20 minutes, to allow the leaves to reach a steady state. Pulses of saturating light (1 second) were applied to measure Fm’ every 60 seconds.
-1 The CO2 concentration was maintained at 390 µL L using an infra-red gas analyser (IRGA;
CIRAS-1, PP systems Ltd, Herts, UK). Fluorescence levels were estimated as described in Figure
2.2 and parameters calculated using the equations below (Maxwell and Johnson, 2000; Golding and
Johnson, 2003).
Fm
Fm'
Ft
Fo SP on AT on SP on
ML on
Figure 2.2. Typical fluorescence signal showing all the reference points. The zero level of
fluorescence (Fo) is measured after measuring light is switched on (ML). Saturating pulse (SP) is
applied and allows to measure maximum fluorescence (Fm). Then actinic light (AT) which drives
photosynthesis is switched on. After 40 seconds, the application of another saturating pulse (SP),
gives the maximum fluorescence in the light (Fm'). Ft is the level of fluorescence immediately
before the saturating pulse (Maxwell and Johnson, 2000) 123 Maximum PSII efficiency, Fv/Fm = (Fm - Fo)/ Fm
Efficiency of Photosystem II (ФPSII) = (Fm' - Ft)/ Fm'
Non-photochemical quenching (NPQ) = (Fm - Fm')/ Fm' (Bilger and Björkman, 1990)
Fv/Fm is a measure which gives the quantum efficiency if all PSII centres were open (Genty et al.,
1989; Maxwell and Johnson, 2000). Changes in this value occur due to changes in the efficiency of
NPQ. Dark-adapted values of Fv/Fm are used as an indicator of plant photosynthetic performance.
In healthy leaves, the optimum Fv/Fm is around 0.83 in most plant species (Björkman and Demmig,
1987; Johnson et al., 1993, Maxwell and Johnson, 2000). ФPSII measures the proportion of the light absorbed by chlorophylls associated with PSII which is then used in photochemistry (Genty et al., 1989; Maxwell and Johnson, 2000). It shows a linear relationship with the efficiency of CO2 fixation under non photorespiratory (low O2) conditions. Under stress conditions, the rate of photorespiration and pseudocyclic electron flow changes. Therefore, the relationship between the efficiency of CO2 fixation and ФPSII changes (Fryer et al., 1998). ФPSII is used to calculate relative linear electron transport of PSII,
Relative electron transport rate of PSII (PSII ETR) = ФPSII x actinic light
To calculate the absolute rate of electron transport through PSII, it is necessary to estimate the absorption of light by the leaf and the proportion of that energy reaching PSII. The former is often assumed to be 0.84 and the latter 0.5, however this is known that in fact both these values vary between species and between plants exposed to different environmental conditions. Estimation of these parameters on a leaf by leaf basis is difficult so, in this thesis, only the relative ETR is estimated.
124 NPQ value is linearly related to the energy dissipated as heat in PSII (Maxwell and Johnson, 2000).
In a typical plant, this value varies from 0.5 to 3.5 at saturating light and depends on the plant species and the on the previous history of the plant. The correct determination of Fo and Fm values is necessary for the quantification of NPQ (Maxwell and Johnson, 2000; Roháček et al., 2008).
3. Measuring P700+ (redox state of P700)
Once the steady-state was achieved, P700 oxidation which is induced by the actinic light was measured during a 100 milliseconds period of darkness. This dark pulse was repeated at 5 seconds intervals, to give an average signal of 20 accumulations. For a given leaf, the signal (P700+) change induced by the light-dark transition was normalized to the corresponding FR-induced signal
(P700total) which gives the proportion of P700 that was oxidized under steady-state conditions and could then be rapidly re-reduced following a transition to darkness. This measure gives the redox state of P700 (Figure 2.3).
+ + Proportion of P700 = P700 /P700total
Decay of P700+ (re-reduction of P700+) was found to approximate to a first-order kinetic, yielding a pseudo-first order rate constant (k) when data were fitted with a single exponential decay equation
-kt (A(t) = A(0) x ℮ + C where, A(t) is the quantity at time t, A(0) is the initial quantity or quantity at time = 0, k is decay constant and C is the constant of integration) (Genty and Harbinson, 1996; Ott et al., 1999; Wientjes and Croce, 2012). The rate of electron transport through PSI was calculated by multiplying the rate constant and the proportion of P700+ (Klughammer and Schreiber, 1994; Ott et al., 1999; Clarke and Johnson, 2001; Golding and Johnson, 2003; Klughammer and Schreiber,
125 2008).
Relative electron transport rate (ETR) of PSI = (proportion of P700+) x k
P700+
+ AT on AT off P700 fraction (P700 ) which is oxidized by actinic light and re- reduced following a transition to darkness P700
Figure 2.3. P700 oxidation of which is induced by the actinic light was measured during a 100 milliseconds period of darkness. The signal (P700+) change induced by the light-dark transition was
+ normalized to the corresponding FR-induced signal (P700total). Decay of P700 (re-reduction of
P700+ during darkness) was found to approximate to a first-order kinetic, yielding a pseudo-first order rate constant (k).
4. Measuring the active P700 pool (P700 active)
The relative concentration of 'active' PSI centres (centres that can be oxidized by light and are then rapidly re-reduced during a period of darkness) was measured (Golding and Johnson, 2003). In addition to the actinic light, 100 milliseconds flashes of 7500 µmol m-2 s-1 were applied to give the maximum P700 signal which represents the fractions of P700+ (oxidized by actinic light) and P700
(open centres). This was followed a transition to darkness. The resulting signal was normalized by
126 the FR induced signal for the same leaf. This measure gives the proportion of 'active' PSI centres
(Figure 2.4) (Klughammer and Schreiber, 1994; Ott et al., 1999; Clarke and Johnson, 2001; Golding and Johnson, 2003; Klughammer and Schreiber, 2008).
Maximum P700 (fractions of AT & SP off P700+ and P700) signal SP on AT on induced by combined actinic illumination plus saturating flash
Dark period
Figure 2.4. The relative concentration of 'active' PSI centres (centres that can be oxidized by light and are then rapidly re-reduced during a period of darkness). In addition to the actinic light (AT),
100 milliseconds flashes of 7500 µmol m-2 s-1 (SP) were applied and this was followed a transition to darkness. This gives the total of the fractions of P700+ and P700. The resulting signal was normalized to FR induced signal for the same leaf.
2.3.3. Chlorophyll content measurements
Leaves of salt-treated and control plants was collected and washed with distilled water. A piece of leaf, approximately 2.1 cm2 (length 3 cm x width 0.7 cm) taken between 4 cm and 7 cm from the leaf tip of each leaf (Figure 2.5) was ground using a mortar and pestle and extracted in a total volume of 10 ml acetone of 80% (v/v). The chlorophyll extraction was centrifuged (3,000 xg) for 5
127 minutes. Chlorophyll content of the supernatant was estimated by measuring absorbance using an
Ocean Optics USB2000 spectrophotometer (Ocean Optics, Dunedin, USA). The following equations were used to convert the absorbance value into chlorophyll content per unit leaf area
(nmol/cm2) (Porra, et al., 1989),
7cm tip base 4cm
Figure 2.5. The first leaf of a barley plant showing the section of approximately 2.1 cm2 (length 3 cm x width 0.7 cm between 4 cm and 7 cm from the leaf tip of each leaf) area which is used for chlorophyll fluorescence, PSI electron transport, gas exchange and chlorophyll content measurements.
Chl a = 13.71 x (A663 - A750) - 2.85 x (A646 - A750)
Chl b = 22.39 x (A646 - A750) - 5.42 X (A663 - A750)
Chl a + Chl b = 19.54 x (A663 - A750) + 8.29 x (A646 - A750)
Where A663 - A750 and A646 - A750 is the difference in absorbance measured at 646 or 663 and 750 nm.
2.3.4. Statistical Analysis
Results are reported as the mean ± standard error of mean (SEM) of at least four replicates from four independent experiments. Significance of results was tested using Analysis of variance
(ANOVA) and Tukey's post-hoc test as indicated in figure legends. Statistical software, SPSS
128 Statistics 20 (IBM) was used for all analysis. P value ≤ 0.05 were considered statistically significant.
129 2.4. Results
Barley plants were treated with 0, 50, 100 and 250 mM NaCl solutions for 14 days before measuring gas exchange parameters, chlorophyll fluorescence, chlorophyll content in leaves and PSI redox state and electron transport rate. Similar to other plant species, tolerance to salt stress may differ with developmental stages in barley (Mano and Takeda, 1997). An early study by Ayers et al. (1952) showed that, compared to the germination stage barley plants are susceptible to salinity during the seedling and early vegetative growth stage. Therefore, this study was performed to examine the effects of salt on the early vegetative stage of barley up to the development of the first 4 leaves. The measurements were taken on the first leaf from four barley plants and measurements were repeated four times for each salt treatment. Effects of salt stress on the germination stage of barley were not tested.
In this experiment, 50 mM is the minimum and 250 mM (equivalent to ~50% sea water) is the maximum salt concentrations used. Although barley is considered as a salt tolerant crop compared, to other crops, such as rice and wheat (Maas, 1990) it is sensitive to salt concentrations higher than
250 mM when exposed for long periods of time (Munns et al., 2006). A study by Pérez-López et al.
(2012), which is in some respects similar to the current study, used 80 mM as the minimum salt concentration and 240 mM as the maximum salt concentration.
Plant responses to salt stress occurred due to both osmotic stress and ionic stress (Munns and Tester,
2008). Plants show rapid responses for osmotic stress whereas, responses occur due to the accumulation of Na+ in leaves are slow and showed when plants are exposed to salt for a long period of time. Barley is a salt includer, where Na+ is translocated to the shoots, rather than be retained in the roots, and it accumulates salt as a tolerance strategy (Glenn et al., 1999; Mian et al.,
130 2011). Therefore in the current study, barley plants were challenged with salt for 14 days to trigger both osmotic and ionic stress responses.
2.4.1. Gas-Exchange Parameters
Gas exchange of control and salt treated barley plants was measured after 14 days of salt treatment.
The rate of assimilation, stomatal conductance and rate of transpiration were measured under 390
-1 µL L CO2 concentration at different light intensities. Control plants showed the maximum rate of assimilation and this decreased with increasing salt concentrations (Figure 2.6). Exposure of plants to salinity induced stomatal closure, which is evident in the decline of transpiration rate. At ambient
CO2 concentration, barley plants treated with 250 mM showed a lower stomatal conductance compared to the control plants. A decrease in the rate of CO2 assimilation, transpiration and stomatal conductance occurred at all salt concentrations at all irradiances, suggesting that the quantum yield of photosynthesis was reduced.
Changes in the rate of CO2 assimilation with varying calculated internal CO2 concentrations were measured at 1000 µmol m-2 s-1 light in control and salt-treated barley plants (Figure 2.6.d). The ACi curve was measured on leaves, with CO2 being decreased and then increased. Despite having considerably lower stomatal conductance, the relationship between assimilation and internal CO2 concentration in plants treated with 50 mM salt concentration was not significantly different to the control. This indicates that the main effect at this salt concentration is to reduce CO2 entry into the plants. However, at higher salt concentrations, including 100 and 250 mM the A/Ci relationship was markedly changed. This shows that, at higher salt concentrations, carbon fixation could not be
-1 restored to control levels by increasing CO2 concentrations even as high as 1600 µL L .
131 a b
c d
Figure 2.6. Gas exchange parameters of barley plants subjected to varying degrees of salinity (a) rate of CO2 assimilation (b) stomatal conductance and (c) rate of transpiration were measured.
Plants were exposed to: 0 (black squares), 50 (red circles), 100 (blue triangles) or 250 (pink down triangles) mM NaCl for 14 days prior to measurements. Leaves were exposed to different actinic
o -1 light for 20 minutes at 25 C in the presence of 390 µL L CO2. (d) Assimilation of CO2 as a
-2 -1 function of internal CO2 concentration (A/Ci curve) was measured at 1000 µmol m s . Symbols as above. The error bars represent the standard error of at least 3 replicates.
132 2.4.2. Chlorophyll Fluorescence Analysis
The maximum quantum efficiency (Fv/Fm), is an indicator of the potential quantum efficiency of
PSII. The optimal mean value of Fv/Fm is around 0.83 in many plants (this value varies with the plant species) and this value decreases when plants are exposed to stress (Björkman and Demming,
1987; Johnson et al., 1993; Maxwell and Johnson, 2000). The maximum quantum efficiency declined with increasing salt concentrations and plants treated with 250 mM salt have the lowest
Fv/Fm compared to the control plants (Figure 2.7).
a a 0.8 b 0.7 c 0.6 0.5 m F
/ 0.4 v F
0.3 0.2 0.1 0.0 0 50 100 250 Salt Concentration (mM)
Figure 2.7. Maximum quantum yield (Fv/Fm) of control and salt-treated barley plants. Plants were treated with 50, 100 or 250 mM NaCl concentrations for 14 days prior to measurements. The error bars represent the standard error of at least 3 replicates. There was a statistically significant difference between Fv/Fm values when exposed to different salt concentrations as determined by one-way ANOVA (p ≤ 0.05). A Tukey's post-hoc test (results are shown as letters above columns) revealed that Fv/Fm value was statistically significantly lower when plants treated with 100 mM and
250 mM NaCl compared with 50 mM. There were no statistically significant differences between the control plants and the plants treated with 50 mM. Mean Fv/Fm values not sharing the same lowercase letter are significantly different.
133 The parameter ФPSII gives a measure of PSII efficiency under any given set of conditions in plants
(Genty et al., 1989). With increasing light intensity, ФPSII declines, reflecting the saturation of PSII photochemistry. With increasing exposure to salt, ФPSII decreased at lower irradiances, showing that PSII photochemistry is more readily saturated (Figure 2.8.a). Control plants showed the highest
ΦPSII, whereas plants treated with 250 mM NaCl showed the lowest. Figure 2.8.b shows the relative electron transport rate of PSII (PSII ETR) of control and salt-treated barley plants with increasing light intensities. With increasing salt stress, PSII ETR decreased. Control plants showed the highest PSII ETR and the plants treated with 250 mM showed the lowest ETR.
Non-photochemical quenching (NPQ) was measured in control and salt-treated plants. In control,
NPQ saturated at a lower value than in the presence of salt. Plants treated with 100 mM showed the maximum NPQ with this reaching a value of 2.5 compared to 2.0 in the control plants. However, plants treated with 250 mM showed low NPQ values at low light intensities compared to other salt- treated plants and a lower saturated levels at high light.
134 a b
c
Figure 2.8. The effect of salt treatment on (a) the efficiency of PSII (ΦPSII) (b) relative electron transport rate of PSII (PSII ETR) and (c) non-photochemical quenching (NPQ) of barley plants.
Plants were exposed to: 0 (black squares), 50 (red circles), 100 (blue triangles) or 250 (pink down triangles) mM NaCl for 14 days prior to measurements. Leaves were exposed to different actinic
o -1 lights for 20 minutes at 25 C in the presence of 390 µL L CO2. The error bars represent the standard error of at least 3 replicates.
135 2.4.3. PSI Photochemistry
In addition to chlorophyll fluorescence, measurements of P700, the primary electron donor of PSI, were performed using absorbance in the near infra-red as an indicator of P700 redox state.
Measurements of the decay of P700 signal following a light- dark transition give information about
PSI electron transport. Under saline conditions, barley showed a higher proportion of PSI reaction centers being oxidized at any given light intensity (Figure 2.9.a). Plants treated with 250 mM showed more than 75% oxidation of P700 at an irradiance of 800 µmol m-2 s-1.
In addition to the relative proportion of oxidized P700, the PSI centres that are active (i.e. where
P700 could be oxidized by a flash of saturating light and then re-reduced in darkness) was measured
(Figure 2.9.b). The relative proportion of active PSI centres increased with increasing salt concentrations. Barley plants treated with 250 mM showed highest proportion of active PSI centres compared to the control plants.
The conductance of the electron transport chain was measured as the rate constant (k) for the decay of oxidized PSI centres when transferred from light to darkness. According to previous findings, this rate constant decreases when plants are subjected to stress (Golding and Johnson, 2003; Stepien and Johnson, 2009). Figure 2.9.c showed that, in this study, the rate constant decreased with increasing NaCl concentrations and the lowest rate constant can be seen at the highest salt concentrations. The rate constant for P700 reduction was largely insensitive to irradiance across the range measured. At the lowest irradiance it was not possible to accurately measure k, due to poor signal: noise ratio.
When considering the electron transport rate through PSI (PSI ETR) (Figure 2.9.d) the highest rate
136 was seen in the control plants. PSI ETR, decreased markedly when plants were treated with salt.
However, PSI ETR did not decrease further when plants are treated with 100 and 250 mM of NaCl.
This results suggested that low salt concentrations reduces PSI ETR but high salt concentrations, which inhibits PSII had no further effect on PSI.
137 a b
c d
Figure 2.9. The effect of salt treatment on (a) redox state of P700 (b) relative proportion of the active PSI centres (c) rate constant for P700 reduction and (d) electron transport rate (ETR) of PSI
(PSI ETR) of barley plants. Plants were exposed to: 0 (black squares), 50 (red circles), 100 (blue triangles) and 250 (pink down triangles) mM NaCl for 14 days prior to measurements. Leaves were
o -1 exposed to different actinic lights for 20 minutes at 25 C in the presence of 390 µL L CO2. The error bars represent the standard error of at least 3 replicates.
138 2.4.4. Effects of salt on chlorophyll content of barley leaves
The chlorophyll content measurements were taken after measuring gas exchange, chlorophyll fluorescence and PSI electron transport. Exposure of barley to salt resulted in a progressive drop in leaf chlorophyll content per leaf area (Figure 2.10.a). Chlorophyll content of leaves treated with 50,
100 and 250 mM NaCl dropped by 15%, 26% and 46%, respectively, relative to the control plants.
Chlorophyll a/b ratio showed a progressive increase when exposed to salt concentrations including
50 and 100 mM. However, the ratio showed a drop when exposed to 250 mM salt concentration.
a b a c b b a c d d
Figure 2.10. The effect of salt treatment on the (a) leaf chlorophyll content per leaf area and (b)
chlorophyll a/b ratio in barley. Plants were treated with 50, 100 and 250 mM NaCl concentrations. 14
days after the salt treatments, measurements of chlorophyll content and ratio were taken. The error
bars represent the standard error of at least 3 replicates. There was a statistically significant
difference between chlorophyll content in leaves when exposed to different salt concentrations as
determined by one-way ANOVA (p ≤ 0.05). A Tukey's post-hoc test (results are shown as letters
above columns) revealed that chlorophyll content and chlorophyll a/b ratio were statistically
significantly changed when plants treated with 50 mM, 100 mM and 250 mM NaCl compared the
control plants. Mean chlorophyll content and chlorophyll a/b values not sharing the same lowercase
letter are significantly different.
139 2.5. Discussion
Although barley is considered as one of the most salt tolerant crops, salinity affects barley production worldwide (Maas and Hoffman, 1977; Shannon, 1984; Martinez-cob et al., 1987;
Colmer et al., 2005). Salt affects plants by reducing water availability and by causing ion imbalance and toxicity (Munns, 2005; Parida and Das, 2005). Studies have shown that the majority of annual crops are tolerant to salinity at the germination stage but are most sensitive during the emergence and the early vegetative stages (Läuchli and Epstein, 1990; Maas and Grattan, 1999; Läuchli and
Grattan, 2007). Consistent with these findings, subjecting barley to different salt concentrations brought about a clear salinity response at the early vegetative stage. Excess salt affects photosynthesis in two ways: (1) low rate of CO2 diffusion (flux) into the leaf, caused by reductions in stomatal and mesophyll conductance (stomatal limitations) and (2) disruption of the metabolic potential for photosynthesis (nonstomatal limitations) (Chaves, 1991; Tezara et al., 1999; Ashraf,
2003; Lawlor and Tezara, 2009; Chaves et al., 2011). Stomatal limitations are considered as the major contributor to inhibition of photosynthesis (Cornic and Briantais, 1991). However, studies have shown that nonstomatal limitations, occurring through direct effect of salt on the photosynthetic apparatus, are responsible for low photosynthetic rates in plants (Ball and Farquhar,
1984; Seeman and Critchley, 1985; Seeman and Sharkey, 1986; Tezara et al, 2002; Chaves et al.,
2009). Evidence for both of these mechanisms was seen in this study.
Previous studies have shown that salt decreases the net photosynthetic rate and stomatal conductance in barley (Tavakkoli et al., 2010; Kalaji et al., 2011; Kalaji et al., 2013). Consistent with these findings, salinity has a major impact on the rate CO2 assimilation, rate of transpiration and stomatal conductance in barley variety Chalice (Figure 2.6.a, b, c). The same effect was observed in a study performed by Pérez-López et al. (2012) who showed that the decrease in the rate of assimilation in barley under ambient and high salt concentration was highly related to the
140 decrease in the stomatal conductance. These results suggest that osmotic stress triggered by low water availability causes stomatal closure and leads to a decrease in CO2 assimilation and stomatal conductance in barley. This was also observed in other plants, including rice (Dionisio-Sese and
Tobita, 2000), soybean (Kao et al., 2003) and sorghum (Netondo et al., 2004). However, Perera et al. (1994) showed that ionic stress occurring in cells, due to the accumulation of Na+ in leaves is responsible for the stomatal closure in Aster tripolium under salt stress. Therefore, response curves between CO2 assimilation (A) and internal CO2 concentration (Ci) were plotted to differentiate the limiting effects on photosynthesis into stomatal and nonstomatal factors (Farquhar and Sharkey,
1982).
Despite having considerably lower stomatal conductance, the relationship between assimilation and calculated internal CO2 concentration in plants treated with 50 mM salt concentration is not significantly different to the control (Figure 2.6.d). This indicates that, at low salt concentrations the only effect of salt on barley plants is limiting the entry of CO2 into the leaves which can be overcome by increasing the external CO2 concentration. Therefore, at 50 mM, the limiting effect on photosynthesis was stomatal. Studies performed by Pérez-López et al. (2008 and 2009) have shown that the rate of assimilation decreased with increasing stress in barley under ambient CO2 and the effect of stress became less severe with increasing CO2 concentration. However, the quantum yield, as indicated, both by the Fv/Fm value (Figure 2.7) and the light response curve (Figure 2.8.a) was affected even at 50 mM, indicating the effect was nonstomatal. At higher salt concentrations the relationship between assimilation and internal CO2 is changed, indicating the occurrence of nonstomatal limitations at 100 and 250 mM. Similar effects were also observed in Hordeum vulgare cv. Chariot under drought (Golding and Johnson, 2003) and in Hordeum vulgare cv. Franklin
(James et al., 2006), Phaseolus vulgaris (Seemann and Critchley, 1985; Brugnoli and Lauteri,
1991), Glycine tomentella (Kao et al., 2003) and Olea europea (Centritto et al., 2003; Loreto et al.,
141 2003) under salinity. A study by Pérez-López et al. (2012) found that elevated CO2 reduced both stomatal and metabolic limitations caused by salt in barley variety Iranis.
The maximum quantum efficiency, Fv/Fm is an important parameter which provides a measure of the rate of linear electron transport (Jamil et al., 2007; Tang et al., 2007; Balouchi, 2010). In healthy leaves, it is close to 0.8 in most of plant species, therefore reduction in Fv/Fm value under stress indicates the occurrence of photoinhibition (Baker and Rosenqvist, 2004; Zlatev, 2009; Vaz and
Sharma, 2011). Studies by Golding and Johnson (2003) and Pérez-López et al. (2012) showed that although ФPSII decreases, under any given condition the maximum quantum efficiency, Fv/Fm does not change in barley when exposed to drought and salt respectively. This indicates that barley shows physiological plasticity which enable it to withstand prolonged exposure to abiotic stress (Pérez-
López et al., 2012). A study by Kalaji et al. (2011) showed that, the Fv/Fm value was unchanged in both salt-sensitive and salt-tolerant barley varieties after 24 hours of salt treatment. However, after 7 days, the Fv/Fm was comparatively low in the salt-sensitive variety. A similar effect was also observed in Arabidopsis thaliana where exposure to salt did not have any immediate effect on
Fv/Fm, however, during the development of salt stress over 14 days, this parameter fell in plants exposed to either 100 or 150 mM NaCl (Stepien and Johnson, 2009). Consistent with that, Fv/Fm declined due to prolonged exposure to NaCl, which suggests that photoinhibition is occurring.
Plants treated with 50 mM showed a slight decrease in Fv/Fm but it is not significant, while plants treated with 100 mM and 250 mM showed a significant decrease compared to the control. This suggests that photoinhibition occurred when plants were exposed to higher salt concentrations.
Stepien and Johnson (2009) showed, in Arabidopsis thaliana, a progressive drop in Fv/Fm values under salt stress, indicating the cumulative damage to PSII reaction centres. Photoinhibition is a stress situation where the rate of photodamage of PSII exceeds the capacity of the repair process
(Aro et al., 1993). Photoinhibition decreases PSII photochemistry and overall plant growth (Aro et
142 al., 1993). This effect is also indicated by the decline in ФPSII when barley plants were exposed to salt. A decrease in ФPSII was also observed in studies performed by Belkhodja et al. (1999) and
Pérez-López et al. (2012) in barley and Moradi and Ismail (2007) in rice under salinity. A drop in
ФPSII in salt-treated barley plants occurred due to the down-regulation of energy transduction from the antenna systems to the reaction centers of PSII under salinity. The decrease in ФPSII and the
Fv/Fm in plants treated with 50 mM of NaCl suggested the occurrence of photoinhibition even at low salt concentrations. However, since PSII efficiency is partly sensitive to CO2, the decrease in
CO2 assimilation may be due to low CO2 in leaves rather than photoinhibition. The linear electron transport rate through PSII is considerably affected by the salt, even at 50 mM. The relative electron transport of PSII (PSII ETR) is lowered in part due to a reduction of the proportion of open PSII centres (qP) (Golding and Johnson, 2003). In addition, NPQ is also contributes to lowering the quantum yield of PSII (Genty et al, 1989; Maxwell and Johnson, 2000).
Studies have shown that there is an increase of NPQ when plants are stressed (Roháček, 2002;
Golding and Johnson, 2003; Redondo-Gómez et al., 2006; Tezara et al., 2008; Ribeiro et al., 2009;
Stepien and Johnson, 2009; Silva et al., 2011; Pérez-López et al., 2012). Consistent with previous findings, NPQ increased gradually when barley plants were challenged with salt (Figure 2.8.c).
NPQ is important for plants to dissipate excess energy absorbed as heat and perform photoprotection in stressed plants (Maxwell and Johnson, 2000). Plants treated with 100 mM showed the highest NPQ values and became stable at higher light intensities. Although NPQ of plants treated with 250 mM is comparatively reduced at low light intensities, it is less saturated at high light. This might be due to the loss of the high energy state quenching component of NPQ (qE) and increases in photoinhibition (qI). Work from Golding and Johnson (2003) showed that, under drought, barley plants obtained higher NPQ values and increased in the proportion of PSI centers which could be oxidized with a saturating flash. These 'active' PSI centers were suggested to be
143 involved in cyclic electron transport chain (CET) which generates a pH gradient required to support qE. This is also observed in Arabidopsis thaliana plants when exposed to salt (Stepien and Johnson,
2009). In this study, we also observed an increase in the proportion of active PSI centres (PSI centres could be oxidized by light and rapidly re-reduced in darkness) when plants were exposed to salt (Figure 2.9.b). The results suggest that the 'additional' PSI centres could be involved in the cyclic electron chain when plants are stressed. Although barley plants treated with 250 mM of NaCl showed an increased level of 'active' PSI centres, NPQ values decreased at low light intensities.
According to results of the proportion of oxidized PSI, it is evident that P700 become more oxidized in the presences of salt (Figure 2.9.a). Barley plants treated with 250 mM showed more than 75% oxidation of P700. Work from Stepien and Johnson (2009) showed that, salt-sensitive
Arabidopsis thaliana plants showed more than 72% oxidation of P700 when exposed to salt while salt-tolerant Thellungiella salsuginea did not show any change in the redox state under any given condition when exposed to salt. The primary electron donor of PSI, P700 becomes oxidized with increasing light and is rapidly re-reduced following a transition to darkness. The decay of P700+ fitted to a mono exponential curve which can be described by a pseudo first order rate constant
(Clarke and Johnson, 2001; Golding and Johnson, 2003). The rate constant provides information about the extent to which electron transport to PSI is down-regulated. Studies have shown that, this rate constant decreases with increasing stress conditions (Clarke and Johnson, 2001; Golding and
Johnson, 2003; Stepien and Johnson, 2009). Consistent with these findings, the rate constant declined with increasing salt concentrations (Figure 2.9.c). The product of the rate constant of the decay and the amount of oxidized P700 gives the rate of electron transport through PSI. The highest
ETR of PSI was observed in control plants. ETR of PSI showed a marked decrease when barley plants were treated with salt (Figure 2.9.d). The results of PSII and PSI photochemistry indicate, that, despite of having an increased level of 'active' PSI centres, cyclic electron flow was either not
144 occurring or occur at very low levels in barley plants at the highest salt concentration.
The chlorophylls and carotenoids in leaves are severely affected by salt (Parida and Das, 2005;
Pinheiro et al., 2008; Li et al., 2010; Yang et al., 2011). A decrease in leaf chlorophyll content under salinity was observed in crops, including sunflower (Ashraf and Sultana, 2000; Akram and Ashraf,
2011), alfalfa (Winicov and Seemann, 1990), wheat (Arfan et al., 2007; Perveen et al., 2010) and castor bean (Pinheiro et al., 2008). Salinity affects the levels of chlorophylls in plants either by inhibiting biosynthesis or increasing degradation (Reddy and Vora, 1985; Fang et al., 1998; Eckardt,
2009). However, the extent of the reduction of chlorophyll content under salinity depends on the salt tolerance of the plant species (Ashraf and Harris, 2013). Morales et al. (1992) found that the total chlorophyll content in the salt tolerant barley variety Igri remain unchanged when plants were subjected to salt (CaCl2 50 mM and NaCl 91.3 mM) and salt sensitive variety Albacete showed a decrease in the chlorophyll content. In this study the total chlorophyll content decreased substantially when plants were subjected to high salt concentrations (Figure 2.10.a). Even at 50 mM the total leaf chlorophyll content decreased by 15% compared to the control plants. At 250 mM most of the leaves showed chlorosis and total chlorophyll content was decreased by 46%.
Therefore, we can suggested that Chalice is a salt-sensitive variety in comparison.
The measure of the chlorophyll a/b ratio indicates the change in composition of the thylakoid membrane and positively correlates with the ratio of PSII reaction centre cores to light harvesting chlorophyll-protein complex (LHCII) (Terashima and Hikosaka, 1995). A study by Djanaguiraman et al. (2006) showed that the degradation of chlorophyll-b is more severe than the chlorophyll-a, resulting in an increase in chlorophyll a/b ratio when plants are exposed to salt. LHCII binds the majority of chlorophyll-b and has a low chlorophyll a/b ratio (1.3-1.4) compared to chlorophyll binding proteins associated with the PSII core (Evans, 1989; Green and Durnford, 1996). Therefore,
145 increases in chlorophyll a/b indicates either loss of light harvesting complexes (LHCs) relative to the reaction centres (RC) in photosystems, or loss of PSII compared to PSI or both (Anderson,
1986). Consistent with that, we observed an increase in chlorophyll a/b ratio when plants were exposed to 50 and 100 mM salt concentrations compared to the control plants. However, plants treated with 250 mM showed a decrease in the chlorophyll a/b ratio compared to the control plants.
With this results we can conclude that barley plants probably showed an increase in chlorophyll a/b ratio at low salt concentrations due to loss of LHC. Chlorophyll a/b ratio decreased at 250 mM of
NaCl, possibly due to damage to or controlled loss of the reaction centres of PSII and/or PSI. A reduction in chlorophyll content and an increase in chlorophyll a/b ratio were also observed in salt- treated spinach leaves (Delfine et al., 1999).
According to results from chlorophyll fluorescence and PSI oxidation, there is evidence for a substantial decrease in PSI ETR compared to PSII ETR when plants are exposed to 50 mM of NaCl, which indicates the possible reduction of cyclic electron flow. However, PSI ETR did not decrease significantly compared PSII ETR which was decreased substantially at 100 and 250 mM of NaCl, which might be due to increased cyclic electron flow. However, the rate of election transport in two photosystems measured per reaction centres and depends on the number of reaction centres (PSI:
PSII) ratio. At 50 mM, despite having low cyclic electron flow plants have higher NPQ, which suggests a possible involvement of the Mehler reaction to maintain a pH gradient across the thylakoid membrane. On the other hand, at 250 mM of NaCl, cyclic electron transport is probably high. However, NPQ drops at this concentration might be due to the increased leakiness of the thylakoid membranes. Studies of Sharkey (2005) and Sharkey and Zhang (2010) have shown that moderately high temperatures (35-45 oC) induces cyclic electron flow and proton leakage through the membranes. Despite having more 'active' PSI centres, chlorophyll a/b ratio decreased at 250 mM suggesting a specific loss of reaction centers in both photosystems or loss of PSI compared to
146 PSII. Therefore, further experiments were needed to analyse the effects of salt on light harvesting complexes and reaction centres in two photosystems. This could be performed by examining the changes in protein contents in these complexes using immunoblot analysis. Apart from that, another study should be performed to analyse the membrane leakiness and this could be measured by using the electrochromic shift (Witt, 1979). This method which uses three wavelengths (505, 520, 535 nm) to exclude interfering signals from light scattering and zeaxanthin, gives the pH component of the proton motive force (Zhang et al., 2009; Sharkey and Zhang, 2010). In addition, relaxation kinetic studies will provide information about the qE and photoinhibition components in NPQ in the salt-treated barley leaves.
Under salt stress, plants are able to protect themselves from destructive ROS by regulating the electron transport chain. Results suggest that at low salt concentrations plants protect PSII centers from excitations pressure by down-regulating the electron transport chain and maintaining a pH gradient across the thylakoid membrane by cyclic electron transport associated with PSI to support
NPQ. However, at the highest concentration of salt examined this regulation starts to fail. The failure might result from a specific loss of PSI, resulting in reduced cyclic electron flow or an increase in the leakiness of the thylakoid membranes resulting in loss of the pH gradient. Salinity causes either short term or long term effects on plant. Short term effects occur within few hours or
1-2 days of the salt treatment, therefore plants show responses due to osmotic stress. On the other hand, long term effects occur after several days of exposure and plants show responses due to both osmotic and ionic stress (Munns and Tester, 2008). Short term measurements provide information of the rapid responses to salinity without changes in protein content in leaves. Long term effects, involving changes in proteins in leaves, will develop over time. For example, effects such as photoinhibition may be progressive or even change within a day. In this study, effects of salinity on barley were shown after 14 days of salt treatment. Because of that, most of the data provide only
147 long term photosynthetic responses of barley to salt rather than both long term and short term salinity responses. Therefore, it is important to extend this study by performing time course experiments showing both short term and long term responses of barley to salt stress. Salt will not however enter leaves immediately, but may change gradually over time, so it is important to monitor leaf salt concentrations at the same time. Apart from that, a comparative study with a salt tolerant barley cultivar (Ligaba and Katsuhara, 2010) or species, such as Hordeum maritimum
(Lombardi et al., 2000) will provide important information on the salt tolerance of barley cultivar,
Chalice.
148 Chapter 03
Physiological evaluation of salinity stress in two rice varieties from Sri Lanka
Sashila Abeykoon Walawwe
Giles N. Johnson
149 Preface
Sashila Abeykoon Walawwe is the primary author of this paper.
Plant growth by Sashila Abeykoon Walawwe
Measurements of chlorophyll content and leaf area by Sashila Abeykoon Walawwe
Photosynthetic measurements of PSII and PSI by Sashila Abeykoon Walawwe
150 3.1. Abstract
The effects of salinity on the regulation of electron transport through photosystem I and photosystem II (PSI and PSII) have been studied in two rice varieties from Sri Lanka. At-354 is a salt-tolerant variety, widely growing in the coastal regions and high saline soils in Sri Lanka. Bg
-352, is a salt-sensitive variety and popular in many regions of the country.
Both varieties of rice at the early vegetative and the flowering stages were treated with 50 and 100 mM NaCl. 14 days after the imposition of salt treatment, the effects of salt stress on gas exchange, chlorophyll fluorescence, PSI electron transport, chlorophyll content and leaf area were examined.
Exposure to salt decreased the rate of CO2 assimilation, stomatal conductance, the rate of transpiration, electron transport rate of PSII (PSII ETR), electron transport rate of PSI (PSI ETR), chlorophyll content and leaf area in both rice varieties. Chlorophyll fluorescence measurements indicated that in both varieties, electron flow through PSII decreased with increasing salt concentration. However, salt treated At-354 showed higher PSII ETR than Bg-352. At higher salt concentrations, low non photochemical quenching (NPQ) in Bg-352 demonstrated that, when plants are stressed, the ability to dissipate excess energy as heat is lowered. PSI was more oxidised in both varieties when exposed to stress. PSI ETR was higher in both varieties when stressed. However, PSI
ETR was lowered in Bg-352 when exposed to the highest salt concentration at the flowering stage.
This might be due to low conductance of the electron transport chain. This effect was not observed in At-354. Salinity lowered the amount of chlorophyll and caused stomatal closure in both varieties but the impact is more pronounced in Bg-352. With these results, it is evident that the regulation of photosynthesis in At-354 is more prominent than in salt-sensitive Bg-352.
151 3.2. Introduction
Rice (Oryza sativa) is the staple crop of many countries around the world, and provides 20% of the daily calorie intake (Negrão et al., 2011). The physical requirements for rice growing limit rice production to certain areas of the world (Food and Agriculture organization of the United Nations
Statistical Yearbook, 2013). Asia is the largest rice producing and consuming region in the world, with more than 150 kg per capita per year rice production (Food and Agriculture organization of the
United Nations Statistical Yearbook, 2013). It is a tropical C3 plant and requires high average temperature during the growing season (Lafitte et al., 2004; Food and Agriculture organization of the United Nations Statistical Yearbook, 2013). Unlike many other plants, rice can grow well in waterlogged or water saturated soils and shows tolerance to submergence. However, it is sensitive to other abiotic stresses, including drought, cold and salinity (Lafitte et al., 2004).
Soil salinity is one of the major problems causing reductions in rice production worldwide (Flowers and Yeo, 1995). Rice is considered as a salt sensitive crop compared to other cereals (Maas and
Hoffman, 1977; Chinnusamy et al., 2005; Munns and Tester, 2008). However, its sensitivity to salt varies with different growth stage. Rice plants are more tolerant during germination and active tillering than during the seedling and reproductive stages (Lutts et al., 1995; Khan et al., 1997;
Shannon et al., 1998; Zeng and Shannon, 2000; Lafitte et al., 2004). Depending on the severity of the stress, physiological parameters such as photosynthesis and plant growth are affected (Negrão et al., 2011). Downregulation of photosynthesis and growth decrease plant yield (Zeng et al., 2002).
Prolonged exposure to salt reduces rice yield by reducing the number of tillers, panicle length, spikelet numbers per panicle, spikelet fertility, grain yield, panicle emergence and flowering
(Khatun and Flowers, 1995; Zeng and Shannon, 2000, Grattan et al., 2002; Gay et al., 2009). Apart from that, it also reduces the percentage of seed set, by reducing pollen viability (Khatun and
Flowers, 1995). A study by Grattan et al. (2002) showed that, if the stress is severe (NaCl > 100
152 mM) rice plants die before maturity, however, even if it is less severe (< 50 mM) the long term exposure will cause delays in panicle initiation and flowering.
High levels of salt cause osmotic stress, due to the high solute concentration in the soil, and ion specific stress, due to changes that occur in the ratios of K+, Na+ and Cl- (Negrão et al., 2011). Salt in soil changes plant water relations, as has been observed in rice plants which show a rapid and temporal drop in stomatal conductance and growth rate as short term responses when exposed to salt concentrations higher than 50 mM (Yeo et al., 1991; Moradi and Ismail, 2007). This is also observed in plants treated with KCl, mannitol or polyethylene-glycol (Yeo et al., 1991; Chazen et al., 1995). Work from Ismail et al. (2007) suggested that a decrease in the stomatal conductance before noticeable changes in the leaf water potential, as a short term stress response to salinity implies an effective communication between root and other parts of the plant. A study by Moradi and Ismail (2007) demonstrated that salt-sensitive genotypes showed a relatively slower reduction in stomatal conductance than salt-tolerant genotypes. Martinez-Atienza et al. (2007) showed the presence of rice orthologs of SOS1, SOS2 and SOS3, which are involved in the SOS pathway of
Na+ control in cells. In addition, the HKT-type transporters (OsHKT1 and OsHKT8) control the entry of Na+ into the roots and maintain low Na+/K+ ratio in cells (Golldack et al., 2003; Ren et al.,
2005). OsHKT1 is involved in removing Na+ from xylem sap in rice plants when exposed to salt
(Ren et al., 2005; Sunarpi et al., 2005; Horie et al., 2006; Davenport et al., 2007; Horie et al., 2009).
Osmotic adjustment through the accumulation of the compatible solutes is one of the long term stress responses in plants (Negrão et al., 2011). Accumulation of the compatible solute proline in the cytoplasm acts against hyperosmotic stress under salinity (Demiral and Türkan, 2006). Studies have also shown that rice plants transformed with genes required for glycine betaine production or trehalose synthesis show an increase in salt tolerance (Garg et al., 2002; Mohanty et al., 2002).
153 Another study has shown that, a transgenic rice over-expressing OsRab7, which encodes a small
GTP (guanosine triphosphate) binding protein, showed enhanced seedling growth and increased proline content under salt-treated conditions (Peng et al., 2014). Intracellular ion compartmentalization is an adaptive mechanism in many plants to survive in saline soils (Negrão et al., 2011). Movement of excess ions, especially Na+, from the cytoplasm to vacuole, is mediated by
Na+/H+ antiporters, which are driven by a proton electrochemical gradient are important in this long term stress response (Blumwald, 1987; Negrão et al., 2011). Overexpression of OsNHX1 (NHX- type antiporter) helped to maintain growth of rice plants under salt stress (Fukuda et al., 2004).
Apart from that, four other NHX-type antiporter genes (OsNHX2 to OsNHX5) were identified in rice and these antiporters were shown to be involved in ion compartmentalization under salt stress
(Yao et al., 2010). Salt tolerant genotypes of rice showed an increase in the activity of ascorbate peroxidase and peroxide dismutase under salinity stress (Moradi and Ismail, 2007). A study by Chen and Gallie (2004) showed an increase in ascorbate redox state involved in raising the stomatal conductance due to increased total open stomata area.
To overcome the problems with soil salinity, new varieties of rice should be introduced through breeding programs (Negrão et al., 2011). Therefore, it is important to understand the physiological, biochemical and genetic control behind the salt tolerance mechanisms in rice. Physiological, biochemical and molecular analysis performed over past few years provided important information on the salt tolerance in rice (Flowers et al., 2000; Zhu, 2001; Zhu, 2002; Xiong et al., 2002;
Flowers, 2004; Chinnusamy et al., 2005; Mahajan and Tuteja, 2005; Munns and Tester, 2008;
Mahajan et al., 2008; Türkan and Demiral, 2009; Negrão et al., 2011; Mohammadi-Nejad et al.,
2010; Horie et al., 2012; Kumar et al., 2013; Mardani et al., 2014). Some of the traditional landraces and cultivars of rice are more salt tolerant than commonly grown cultivars (Walia et al., 2005).
These traditional salt tolerant cultivars are good sources of salt tolerant traits, therefore, they are
154 widely used in salt tolerance studies and breeding programs. However, these traditional cultivars have poor agronomic traits, including low yield, poor grain quality and photosensitivity. Pokkali is one such salt tolerant traditional cultivar, popular in South Asia and intensively used in breeding programs to produce salt tolerant varieties (Walia et al., 2005). In this study, we aimed to improve our knowledge of the responses of the rice photosynthetic apparatus to salinity stress through using two rice cultivars from Sri Lanka. Bg-352 is a salt-sensitive, commonly grown cultivar and At-354 is a salt-tolerant cultivar. The knowledge gained from this study can use for future breeding programs to produce high yielding salt tolerant rice varieties.
Bg-352 (common name White Nadu) is one of the commonly grown hybrid rice varieties, which is sensitive to salinity. However, it is resistant to blast, brown plant hoppers and gold midge attacks.
The life span is about 3.5 months and the estimated yield is 10 tons per hectare (Sirisena et al.,
2010). Bg-352 is popular among farmers because of high yield and wide adaptability (Jayawardene,
2011). At-354 (common name Red Nadu) is a salt tolerant variety, developed from a cross between
Pokkali and the high-yielding and erect-leaved variety Bg-94-1 (De Costa et al., 2012). This variety is widely grown in the coastal areas of Sri Lanka (Sirisena et al., 2010). The life span of this rice variety is also 3.5 months and it forms an erect-leaved canopy (De Costa et al., 2012). The estimated yield under optimal growth conditions is about 10 tons per hectare and 5 tons per hectare under saline conditions (De Costa et al., 2012). Compared to Pokkali, At-354 shows a slower rate of
Na+ accumulation in shoots, therefore has lower rates of reduction in relative plant biomass and relative leaf area in response to salt (De Costa et al., 2012). Although, a little research work has been carried out to characterise features of these two rice varieties, no recorded data have focussed on understanding the physiology of photosynthesis in these and characterizing the salt tolerant traits in rice which are responsible for the regulation of photosynthesis. Therefore, the aim of this study was to understand the effects of salinity on photosynthesis in these two rice varieties at the early
155 vegetative stage (tillering) and the flowering stage (reproductive) by measuring gas-exchange, chlorophyll fluorescence, PSI electron transport, leaf area and chlorophyll content.
156 3.3.Materials and Methods
3.3.1. Plant growth and Treatment
Rice seeds (Oryza sativa, varieties, At-354 and Bg-352) were obtained from and certified by the
Rice Research and Development Institute (RRDI) in Batalagoda, Sri Lanka.
1. Plants at the early vegetative stage
Plants were grown in a controlled environment room with a photon flux density of 280 µmol m-2 s-1 provided by white fluorescent bulbs on a 16 hour light/8 hour dark cycle. The day time temperature was 25 oC and night time temperature was 15 oC. Seeds were germinated and were grown in pots filled with John Innes No 3 soil for 20 days and then plants were treated with 100 ml of NaCl solutions of 50 mM or 100 mM concentrations for 14 days (treated 3 times per week). The first leaf of four rice plants in the same pot were taken for the measurements. Measurements for gas- exchange, chlorophyll fluorescence, PSI electron transport, chlorophyll content and leaf area were repeated on 4 rice plant pots for each salt treatment and measurements being performed on the leaf between 5 and 8 cm from the leaf tip.
2. Plants at the flowering stage
Plants were also grown in a greenhouse heated at 30 oC at the botanical grounds of the University of
Manchester. Seeds were germinated and grown in pots containing mixture of soil (50% grit and
50% John Innes No 3 soil based compost) for two months and treated with 100 ml of NaCl solutions of 50 mM or 100 mM concentrations for 14 days (treated 3 times per week). The first leaf around the panicle from two rice plants were used for the measurements. Measurements for gas-exchange,
157 chlorophyll fluorescence, PSI electron transport, chlorophyll content and leaf area were repeated on
4 rice plant pots for each salt treatment and measurements being performed on the leaf between 5 and 8 cm from the leaf tip.
3.3.2. Measuring Photosynthetic parameters (Gas exchange, P700 oxidation and chlorophyll fluorescence measurements)
3.3.2.1. Measuring gas exchange
The measurements of gas exchange were performed in combination with chlorophyll fluorescence analysis. Gas exchange measurements were taken using an infra-red gas analyser (IRGA; CIRAS-1,
PP systems Ltd, Hitchin, UK). Leaves of rice plants were clamped in the cuvette chamber to fill the chamber without overlapping. Temperature and humidity were maintained at 25 oC and 60%, respectively. Leaves were left to equilibrate in the dark for 5 minutes. The rate of gas exchange was measured for 1-2 minutes. After that, leaves were illuminated for 20 minutes until a stable rate of gas exchange was obtained (Johnson and Murchie, 2011). The external CO2 concentration was maintained at 390 µL L-1 and assimilation rate (A), stomatal conductance to water vapour (gs) and transpiration rate (T) were measured for each actinic light intensity. Actinic light (up to 2,500 µmol m-2 s-1) was supplied by a Luxeon III red LED in a laboratory built lamp and light intensity was measured using a PAR meter (SKP215; Skye Instruments, Powys, UK).
A gas with known concentration of CO2 and water vapour was passed through the chamber at a constant 200 cm3/minute rate. Differences in absorbance in the infra-red (IR) analyser were used to
measure the amount of CO2 and water in gas leaving the chamber, compared to that entering. The flux in CO2 and water per unit leaf area were calculated by measuring the differences in gas
158 concentration between the reference line and the leaf sample leaving the cuvette chamber. The humidity in the internal leaf air spaces, the leaf temperature and the external leaf humidity were used to calculate the total conductance of the leaf to water vapour. This is calculated using the Fick's law of diffusion (Caemmerer and Farquhar, 1981). The stomatal conductance to water vapour was calculated by removing the boundary layer contribution (as estimated by the manufacturer for a given chamber). The CO2 conductance is obtained from the stomatal conductance by correcting for the physical properties of CO2 and water molecules. Internal CO2 concentration, Ci, is the sub- stomatal or mesophyll cell wall CO2 concentration. Ci, is assumed to be the in vivo substrate concentration for Rubisco, calculated using the CO2 conductance, assimilation rate and transpiration rate (Caemmerer and Farquhar, 1981; Johnson and Murchie, 2011).
An ACi curve can be to separate the various limiting steps in photosynthesis, including Rubisco activity, RuBP regeneration, triose phosphate utilization and stomatal limitations (Farquhar and
Sharkey, 1982; Johnson and Murchie, 2011). Measurements of ACi curves were taken under a saturating actinic light intensity of 1000 µmol m-2 s-1. First, leaves in the cuvette chamber was
-1 equilibrated with ambient CO2 level (390 µL L CO2). Then leaves were exposed to series of CO2 concentrations, starting from low values (typically minimum of between 10 and 40 µL L-1). The time lag between two measurements at different CO2 concentrations was restricted to 4 minutes and each curve was completed within 20-30 minutes (Johnson and Murchie, 2011; Pérez-López et al.,
2012). CIRAS-1 software was programmed to calculated Ci value for each CO2 concentrations used.
3.3.2.2. Chlorophyll Fluorescence and PSI measurements
Changes in absorbance at 830-870 nm were used to give a measure of the redox state of the PSI
159 primary donor, P700 (Harbinson and Woodward, 1987; Klughammer and Schreiber, 1994).
Measurements were made using a Walz PAM 101 fluorometer in combination with an ED-
P700DW-E emitter-detector unit (Walz, Effeltrich, Germany). Actinic light (up to 2, 500 µmol m-2 s-
1) was supplied by a Luxeon III red LED in a laboratory built lamp. The data were observed and captured using a National Instruments PCI-6220 data acquisition card, in a computer running software written using Labview (National Instruments, Austin, TX, USA). Chlorophyll fluorescence measurements of PSII were made using a PAM 101 fluorometer together with a 101-ED emitter- detector unit (Walz). Fluorescence data were recorded and captured using the same software
(Golding and Johnson, 2003; Hald et al., 2008).
For measurements on any given leaf sample, the following sequence of procedures were carried out:
1. Maximum P700 signal
To measure the signal corresponding to 100% of P700, dark-adapted leaves were exposed to far-red light, maximum 730 nm provided by a far red LED array (Roithner Lasertechnik, Vienna, Austria) for 1 minute. To check whether the light was saturating, a 100 milliseconds flash of red light (4000
µmol m-2 s-1) was applied on top of the far-red light. The saturation of the far-red light was indicated by the absence of a rise in the signal after applying the flash (Golding and Johnson, 2003). This gives the maximum oxidation of P700 (P700total) (Figure 3.1).
160 SP on Signal size for P700total
FR on
Figure 3.1. Far-red light (FR) induced signal giving 100% of P700. Leaves were exposed to far-red light, λmax = 730 nm, for 1 minute. A 100 milliseconds flash of red light (SP) (4000 µmol m-2 s-1) was applied on top of the far-red light to check whether P700 was fully oxidised. The absence of a rise in the signal after applying the red flash indicates that the P700 signal was saturated.
2. Chlorophyll fluorescence
Leaves were allowed to recover for two minutes after illuminating with FR light (to re-reduce all the P700+). Fo (zero level of fluorescence) is the dark-adapted initial minimum fluorescence and was measured by applying the PAM-101 measuring light for 10-20 seconds. A high intensity short duration light (1 second pulse of red light with a PFD of 4000 µmol m-2 s-1) was applied to reached the maximum fluorescence level, Fm (maximum fluorescence). Following that, actinic light was switched on and plants were left for 20 minutes, to allow the leaves to reach a steady state. Pulses of saturating light (1 second) were applied to measure Fm' every 50 seconds. The CO2 concentration was maintained at 390 µL L-1 using an infra-red gas analyser (IRGA; CIRAS-1, PP systems Ltd,
Herts, UK). Fluorescence levels were estimated as described in Figure 3.2 and parameters
161 calculated using the equations below (Maxwell and Johnson, 2000; Golding and Johnson, 2003).
Fm
Fm'
Ft
Fo SP on AT on SP on
ML on
Figure 3.2. Typical fluorescence signal showing all the reference points. The zero level of fluorescence (Fo) is measured after measuring light (ML) is switched on. Saturating pulse (SP) is applied and allows to measure maximum fluorescence (Fm). Then actinic light (AT) which drives photosynthesis is switched on. After 50 seconds, the application of another saturating pulse (SP), gives the maximum fluorescence in the light (Fm'). Ft is the level of fluorescence immediately before the saturating pulse (Maxwell and Johnson, 2000).
Maximum PSII efficiency, Fv/Fm = (Fm - Fo)/ Fm
Efficiency of Photosystem II (ФPSII) = (Fm' - Ft)/ Fm'
Non-photochemical quenching (NPQ) = (Fm - Fm')/ Fm' (Bilger and Björkman, 1990)
Fv/Fm is a measure which gives the quantum efficiency if all PSII centres were open (Genty et al.,
1989; Maxwell and Johnson, 2000). A change in this value occurs due to changes in the efficiency of light capture. Dark-adapted values of Fv/Fm can be used as an indicator of plant photosynthetic performance. In healthy leaves, the optimum Fv/Fm is around 0.83 in most plant species (Björkman
162 and Demmig, 1987; Johnson et al., 1993, Maxwell and Johnson, 2000). ФPSII measures the proportion of the light absorbed by chlorophylls associated with PSII which is then used in photochemistry (Genty et al., 1989; Maxwell and Johnson, 2000). It typically shows a linear relationship with the efficiency of CO2 fixation under non photorespiratory (low O2) conditions.
Under stress conditions, the rate of photorespiration and pseudocyclic electron flow changes.
Therefore, the relationship between the efficiency of CO2 fixation and ФPSII changes (Fryer et al.,
1998). ФPSII is used to calculate relative linear electron transport of PSII,
Relative electron transport rate of PSII (PSII ETR) = ФPSII x actinic light
To calculate the absolute rate of electron transport through PSII, it is necessary to estimate the absorption of light by the leaf and the proportion of that energy reaching PSII. The former is often assumed to be 0.84 and the latter 0.5, however this is known that in fact both these values vary between species and between plants exposed to different environmental conditions. Estimation of these parameters on a leaf by leaf basis is difficult so, in this thesis, only the relative ETR is estimated.
3. Measuring P700+ (redox state of P700)
Once a steady-state was achieved, P700 oxidation induced by the actinic light was measured during a 100 milliseconds period of darkness (Figure 3.3). This dark pulse was repeated at 5 seconds intervals, to give an average signal of 20 accumulations. For a given leaf, the P700+ signal change induced by the light-dark transition was normalized to the corresponding FR-induced signal
(P700total) (Figure 3.1) to give the proportion of P700 that was oxidized under steady-state conditions and could then be rapidly re-reduced following a transition to darkness. This measure
163 gives the redox state of P700.
+ + Proportion of P700 = P700 /P700total
Decay of P700+ (re-reduction of P700+) was found to approximate to a first-order kinetic, yielding a pseudo-first order rate constant (k) when data were fitted with a single exponential decay equation
-kt (A(t) = A(0) x ℮ + C where, A(t) is the quantity at time t, A(0) is the initial quantity or quantity at time = 0, k is decay constant and C is the constant of integration) (Genty and Harbinson, 1996; Ott et al., 1999; Wientjes and Croce, 2012). The rate of electron transport through PSI was calculated by multiplying the rate constant and the proportion of P700+ (Klughammer and Schreiber, 1994; Ott et al., 1999; Clarke and Johnson, 2001; Golding and Johnson, 2003; Klughammer and Schreiber,
2008).
Relative electron transport rate of PSI (PSI ETR) = (proportion of P700+) x k
P700+
+ AT on AT off P700 fraction (P700 ) which is oxidized by actinic light and re- reduced following a transition P700 to darkness
Figure 3.3. P700 oxidation induced by the actinic light was measured during a 100 milliseconds period of darkness. The signal (P700+) change induced by the light-dark transition was normalized to the corresponding FR-induced signal (P700total). Decay of P700+ (re-reduction of P700+ during darkness) was found to approximate to a first-order kinetic, yielding a pseudo-first order rate constant (k). 164 4. Measuring the active P700 pool (P700 active)
The relative concentration of 'active' PSI centres (centres that can be oxidized by light and are then rapidly re-reduced during a period of darkness) was measured (Golding and Johnson, 2003). In addition to the actinic light, 100 milliseconds flashes of 7500 µmol m-2 s-1 were applied to give the maximum P700 signal which represents the fractions of P700+ (oxidized by actinic light) and P700
(open centres). This was followed a transition to darkness. The resulting signal was normalized by the FR induced signal for the same leaf. This measure gives the proportion of 'active' PSI centres
(Figure 3.4) (Klughammer and Schreiber, 1994; Ott et al., 1999; Clarke and Johnson, 2001; Golding and Johnson, 2003; Klughammer and Schreiber, 2008).
Maximum P700 (fractions of AT & SP off P700+ and P700) signal SP on AT on induced by combined actinic illumination plus saturating flash
Dark period
Figure 3.4. The relative concentration of 'active' PSI centres (centres that can be oxidized by light and are then rapidly re-reduced during a period of darkness). In addition to the actinic light (AT),
100 milliseconds flashes of 7500 µmol m-2 s-1 (SP) were applied and this was followed a transition to darkness. This gives the total of the fractions of P700+ and P700. The resulting signal was normalized to FR induced signal for the same leaf.
165 5. Relaxation analysis to distinguished slow and rapid relaxation kinetics of NPQ
NPQ is linearly related to the energy dissipated as heat in PSII (Maxwell and Johnson, 2000). In a typical plant, this value varies from 0.5 to 3.5 at saturating light and depends on the plant species and on the previous history of the plant. Changes in NPQ values represent a change in the efficiency of energy dissipation and this is measured relative to the dark-adapted state. Increase in NPQ could be due to photoprotective processes (state transitions, qT and high energy state quenching, qE) or the damage itself (photoinhibition, qI). Therefore, it is important to measure the relaxation kinetics to distinguish each process. After light is removed qE relaxes within seconds to minutes and qT relaxes within tens of minutes. However, these two processes cannot easily be distinguished from their relaxation kinetics (Walters and Horton, 1991). qI relaxes over long period of time (hours)
(Walters and Horton, 1991; Maxwell and Johnson, 2000).
A relaxation analysis was performed to measure slow and rapid relaxation components of NPQ. In this analysis, quenching is allowed to relax and the maximum fluorescence levels (Fm'') were recorded at regular intervals (Figure 3.5) (Maxwell and Johnson, 2000). The actinic light was switched off and a high intensity short duration light (1 second pulse of red light with a PFD of
4000 µmol m-2 s-1) was applied at 3 minutes intervals over 45 minutes. Fm'' was recorded after applying each saturating pulse. The rising values of the saturating pulses after switching off the actinic light, showed the relaxation of NPQ over time. The NPQ relaxation kinetics show an exponential character (Roháček et al., 2008).
166 Fm
Fm''
Fm'
Fo AT off AT on SP on SP on
Light period Dark period
Figure 3.5. Fluorescence signal showing the relaxation kinetics. The zero level of fluorescence (Fo) is measured after measuring light is switched on. Saturating pulse (SP) is applied and allows estimation of maximum fluorescence (Fm). Then actinic light (AT), which drives photosynthesis is switched on. After 50 seconds, the application of another saturating pulse (SP), gives the maximum fluorescence in the light (Fm'). After the actinic light was switched off and a high intensity short duration light (1 second pulse of red light with a PFD of 4000 µmol m-2 s-1) was applied at 3 minutes intervals for over 45 minutes. Fm'' was recorded after applying the saturating pulse (SP)
A graph was plotted from log(Fm'') against time. The recorded data points were extrapolated toward the point where the actinic light was switched off, gives Fmr. This value was used to calculate the slow and fast relaxation components of unrelaxed NPQ. Followings are the equations used to calculated slow (NPQS) and fast (NPQF) relaxing quenching (Maxwell and Johnson, 2000):
r r NPQS = (Fm – Fm )/ Fm
r NPQF = (Fm/Fm') – (Fm/Fm )
167 3.3.3. Leaf area and chlorophyll content measurements
Leaves of salt-treated and control plants were collected and washed with distilled water. Then the total leaf area was measured using ImageJ software version 1.46. A piece of leaf, approximately 2.4 cm2 (length 3 cm x width 0.8 cm) was taken between 8 cm and 5 cm from the leaf tip of each leaf was ground using a mortar and pestle and extracted in a total volume of 10 ml 80% (v/v) acetone.
The chlorophyll extract was centrifuged (3,000 xg) for 5 minutes. Chlorophyll content of the supernatant was estimated by measuring absorbance using an Ocean Optics USB2000 spectrophotometer (Ocean Optics, Dunedin, USA). The following equations were used to convert the absorbance value into chlorophyll content per unit leaf area (nmol/cm2) (Porra, et al., 1989),
Chl a = 13.71 x (A663 - A750) - 2.85 x (A646 - A750)
Chl b = 22.39 x (A646 - A750) - 5.42 X (A663 - A750)
Chl a + Chl b = 19.54 x (A663 - A750) + 8.29 x (A646 - A750)
Where A663 - A750 and A646 - A750 is the difference in absorbance measured at 646 or 663 and 750 nm.
3.3.4. Statistical Analysis
Results are reported as the mean ± standard error of mean (SEM) of at least four replicates from three independent experiments. Significance of results was tested using Analysis of variance
(ANOVA) and Tukey's post-hoc test as indicated in figure legends. Statistical software, SPSS
Statistics 20 (IBM) was used for all analysis. P value ≤ 0.05 were considered statistically significant.
168 3.4. Results
Rice plants (varieties At-354 and Bg-352) at the early vegetative stage (tillering) and the flowering stage (reproductive) were treated with 50 or 100 mM salt concentrations for 14 days. Salinity affects all stages of growth and development in rice (Flowers & Yeo, 1981; Lutts et al., 1995;
Shereen et al., 2005; Walia et al., 2005). However, sensitivity of rice to salt depends on the growth stage, concentration of the salt solution and the duration of the exposure. Studies of Flowers and
Yeo (1981) and Lutts et al. (1995) showed that rice plants are more sensitive to salt at the seedling stage than the reproductive stage. However, several studies have shown contrasting results (Heenan et al., 1988; Hoshikawa, 1989; Khan et al., 1997; Zeng and Shannon, 2000). Therefore, the current study was focused on examining the effects of salt on two growth stages of rice, the early vegetative and the flowering stages.
50 mM of NaCl is the minimum salt concentration which trigged the salt-induced responses in the salt-tolerant At-354. 100 mM of NaCl is the maximum salt concentration used, where the salt- sensitive Bg-352 showed salinity responses without dying. Plant responses to salt stress occurred due to both osmotic stress and ionic stress (Munns and Tester, 2008). Plants show rapid responses to osmotic stress, whereas responses due to the accumulation of Na+ in leaves are slow and only seen when plants are exposed to salt for a long period of time. Therefore, rice plants were treated with salt for 14 days to trigger both osmotic and ionic stress responses.
3.4.1. Rice plants exposed to different salt concentrations of 50 and 100 mM
Salt stress significantly reduced the efficiency of photosynthesis in two rice genotypes during both the early vegetative and the flowering stage. Control plants of both varieties (34 days old) at the
169 early vegetative stage had leaves which were 20-25 cm long and 0.5-0.7 cm wide (Figure 3.6.a).
Plants showed poor growth and leaves were wilted and showed a decrease in leaf length and width and chlorosis, after the salt treatment (Figure 3.6.b and 3.6.c).
Figure 3.6. 34 days old At-354 and Bg-352 control plants at the early vegetative stage (a). Both varieties of rice plants were treated with 50 and 100 mM of NaCl for 14 days. At-345 plants showed decrease in plant height and leaf size when treated with 50 and 100 mM NaCl for 14 days (b). Bg-
352 plants showed decrease in plant height and leaf size when treated with 50 and 100 mM NaCl for 14 days (c).
170 3.4.2. Effects of salt on leaf area
The total leaf area of the two rice varieties was significantly decreased with increasing salt concentrations. However, the salt-sensitive Bg-352 showed a more prominent reduction in the leaf area than the salt-tolerant At-354 when exposed to salt. At the flowering stage, the first leaf around the panicle was used to measure changes in the leaf area. Compared to the flowering stage, reduction of the leaf area after salt treatments in more prominent in the early vegetative stage
(Figure 3.7).
Early vegetative stage Flowering stage a b
a a a b c d d e b
c
d e
Figure 3.7. Change in leaf area of two rice varieties At-354 (hatched bars) and Bg-352 (white bars)
at the early vegetative stage (a) and at the flowering stage (b) when exposed to 50 and 100 mM of
NaCl. Leaves were collected after 14 days of salt treatment. The error bars represent the standard
error of at least 3 replicates. There was a statistically significant difference between leaf area when
exposed to different salt concentrations as determined by one-way ANOVA (p ≤ 0.05). Tukey's post-
hoc test results are shown as letters above columns. Mean leaf area values not sharing the same
lowercase letter are significantly different.
171 3.4.3. Chlorophyll content measurements
Leaf chlorophyll content and chlorophyll a/b ratio were measured in control and salt treated rice plants. In control plants, leaf chlorophyll content in At-354 and Bg-352 was similar (Figure 3.8).
Upon exposure to salt, the chlorophyll content in both varieties decreased significantly. However,
Bg-352 showed a larger decrease in chlorophyll concentration compared to At-354.
Chlorophyll a/b ratio was similar in both varieties under control conditions. Compared to the control plants, chlorophyll a/b ratio was significantly increased in At-354 in both the early vegetative and the flowering stages, when treated with 50 mM NaCl. Bg-352, on the other hand showed a significant decrease in this ratio when exposed to 50 mM NaCl. Chlorophyll a/b ratio was decreased in both varieties when plants were exposed to 100 mM.
172 Early vegetative stage Flowering stage a b
a a a a b b d d c c e e
c d b b a a a a d c d c e e
Figure 3.8. The effect of salinity on leaf chlorophyll content per leaf area (a, b) and chlorophyll a/b ratio (c, d) in two varieties of rice, At-354 (hatched bars) and Bg-352 (white bars). The early vegetative stage (a, c) and the flowering stage (b, d) were exposed to 50 and 100 mM NaCl concentrations. Leaves were collected after 14 days of salt treatments. The error bars represent the standard error of at least 3 replicates. There was a statistically significant difference between leaf chlorophyll content and chlorophyll a/b ratio when exposed to different salt concentrations as determined by one-way ANOVA (p ≤ 0.05). Tukey's post-hoc test results are shown as letters above columns. Mean chlorophyll content and chlorophyll a/b values not sharing the same lowercase letter are significantly different.
173 3.4.4. Gas Exchange
The rate of photosynthesis, measured as the CO2 assimilation was higher, in control plants of the two rice varieties examined than in the salt-treated plants in both early vegetative and flowering stages (Figure 3.9 a, b). The rate decreased with increasing salt concentrations. In both stages, the
Bg-352 variety showed the lowest assimilation when exposed to salt. Stomatal conductance to water vapour (Figure 3.9 c, d) and transpiration rate (Figure 3.9 e, f) in both rice varieties decreased with increasing salt concentrations. However, Bg-352 showed a more substantial drop in both parameters when exposed to salt than At-354. This decreased with increasing salt concentrations.
The CO2 assimilation rate as a function of internal CO2 concentration (ACi curve) can be used to separate various limiting steps in photosynthesis including regeneration of RuBP, mesophyll conductance and stomatal limitations (Johnson and Murchie, 2011). ACi curves show that the decrease in the assimilation rate in both rice varieties under salt stress may be due to both stomatal and non-stomatal limitations. Both At-354 and Bg-352 showed a decrease in assimilation when exposed to salt, compared to the control plants. However, in Bg-352, the decrease in relationship between assimilation and internal CO2 concentration is more dramatic than in At-354 (Figure 3.10).
This indicates that, in At-354, the decrease in assimilation under salt stress is largely due to stomatal closure rather than non stomatal limitations. The decrease in the assimilation in Bg-352 under high salt concentrations is due to processes not dependant upon the maintenance of Ci, and plants biochemistry is directly affected by salt.
174 Early vegetative stage Flowering stage
a b
c d
e f
Figure 3.9. Gas exchange parameters measured in rice plants at the early vegetative stage (a, c, e) and the flowering stage (b, d, f) of Bg-352 (closed symbols) and At-354 (open symbols) exposed to:
0 (black squares), 50 (blue triangles) and 100 (red diamonds) mM of NaCl. CO2 assimilation rate
(A) (a, b) stomatal conductance (gs) (c, d) and transpiration rate (T) (e, f) were measured. Leaves
o -1 were exposed to different actinic lights for 20 minutes at 25 C in the presence of 390 μL L CO2.
The error bars represent the standard error of at least 3 replicates.
175 Early vegetative stage Flowering stage a b
Figure 3.10. CO2 assimilation rate (A) as a function of internal CO2 concentration (Ci) in
plants at the early vegetative stage (a) and flowering stage (b) of Bg-352 (closed symbols)
and At-354 (open symbols) exposed to: 0 (black squares), 50 (blue triangles) and 100 (red
diamonds) mM of NaCl. Leaves were exposed to different actinic lights for 20 minutes at 25
oC in the presence of 390 μL L-1 CO2. The error bars represent the standard error of at least 3
replicates.
176 3.4.5. Chlorophyll Fluorescence
Measurements of chlorophyll fluorescence provide information about PSII activity in intact plants.
The ratio Fv/Fm, measures the maximum quantum yield of photosystem II. Changes in this value occur due to changes in the efficiency of light capture, caused e.g. by photoinhibition. Low values can been seen when plants are exposed to stress (Maxwell and Johnson, 2000). In control plants in both the early vegetative stage and the flowering stage, the Fv/Fm ratio, is close to 0.8 (Figure 3.11).
Fv/Fm was sensitive to salt treatments in both At-354 and Bg-352 and showed a significant decrease, however the effect of salt treatment was less marked in the salt-tolerant At-354 than in salt sensitive Bg-352 line.
Early vegetative stage Flowering stage a b a a b a a d b c d c e e
Figure 3.11. Maximum quantum yield (Fv/Fm) of salt stressed (50 or 100 mM) and control plants
(a) plants at early vegetative stage (b) plants at flowering stage of At-354 (hatched bars) and Bg-
352 (white bars). The error bars represent the standard error of at least 3 replicates. There was a
statistically significant difference between Fv/Fm value when exposed to different salt
concentrations as determined by one-way ANOVA (p ≤ 0.05). Tukey's post-hoc test results are
shown as letters above columns. Mean Fv/Fm values not sharing the same lowercase letter are
significantly different.
177 The parameter ΦPSII gives a measure of PSII efficiency under any given set of conditions in plants
(Maxwell and Johnson, 2000). With increasing light intensity, ΦPSII declines, reflecting the saturation of PSII photochemistry. In control plants, ΦPSII was not significantly different between the two genotypes at any irradiance. Both genotypes showed a significant decrease in ΦPSII at any given light when plants were exposed to salt (Figure 3.12). The salt sensitive Bg-352 showed a decrease of ΦPSII when treated with high salt concentrations. When plants were treated with
100mM NaCl, both At-354 and Bg-352 showed a drastic decrease in the efficiency.
The relative linear electron transport rate in PSII (PSII ETR) can be estimated by multiplying the efficiency of PSII (ΦPSII) with the light intensity (Genty et al., 1989). PSII ETR increases with light intensity and then saturates. At-354 and Bg-352 control plants showed higher electron transport rates than salt treated plants. PSII ETR of both control and salt-treated plants is higher in
At-354 than Bg-352.
178 a Early vegetative stage Flowering stage b
d c
Figure 3.12. Photochemical efficiency (ΦPSII) (a, b) and relative linear electron transport rate of
PSII (PSII ETR) (c, d) plants at the early vegetative stage (a, c) and the flowering stage (b, d) of Bg-
352 (closed symbols) and At-354 (open symbols) exposed to: 0 (black squares), 50 (blue triangles) and 100 (red diamonds) mM of NaCl. Leaves were exposed to different actinic light for 20 minutes
o -1 at 25 C in the presence of 390 μL L CO2. The error bars represent the standard error of at least 3 replicates.
179 By dissipating excess light energy as heat, plants are able to protect themselves from high light.
This is measured as non-photochemical quenching (NPQ) of chlorophyll fluorescence (Maxwell and
Johnson, 2000). Subjecting rice to a range of NaCl concentrations induced substantial increases in
NPQ at low light (Figure 3.13). NPQ of control plants at the early vegetative stage is low compared to the salt treated plants. Bg-352 treated with 50 mM NaCl showed the highest NPQ which decreased when plants were subjected to 100 mM salt, showing that the salt sensitive Bg-352 plants lack the ability to dissipate excess energy when plants are stressed with higher salt concentration. At the flowering stage, control plants showed the lowest NPQ value with increasing light intensities. In both varieties, NPQ value increased with increasing salt concentrations. In both the early vegetative and the flowering stages, NPQ of At-354 treated with 100 mM NaCl was not inhibited, whereas Bg-
352 showed a marked decrease when treated with 100 mM NaCl at the early vegetative stage and showed substantial increase in NPQ at the flowering stage.
The increase in NPQ might be due to two processes: protective high-energy-state quenching or photoinhibition. These two processes can be partially distinguished through analysing relaxation kinetics (Maxwell and Johnson, 2000). Both rice varieties showed an increase in both fast and slow- relaxing components of NPQ (NPQF and NPQS) when exposed to NaCl (Table 3.1). The majority of quenching relaxed rapidly in the dark, showing that it was high energy-state quenching (qE) or state transitions (qT) rather than photoinhibition (qI) even at highest salt concentration. However, in both growth stages, the slow-relaxing component of Bg-352 treated with 100 mM is higher than in At-
354. Changes in relaxation kinetics between controlled and salt-treated plants do not impact on the deconvolution used here. Figure 3.14 showed the deconvolution signals of control plants and plants exposed to 100 mM NaCl of both At-354 and Bg-352 at the flowering stage.
180 Early vegetative stage Flowering stage a b
Figure 3.13. Non Photochemical Quenching (NPQ) plants at the early vegetative stage (a) and the flowering stage (b) of Bg-352 (closed symbols) and At-354 (open symbols) exposed to: 0 (black squares), 50 (blue triangles) and 100 (red diamonds) mM of NaCl. Leaves were exposed to different
o -1 actinic lights for 20 minutes at 25 C in the presence of 390 μL L CO2. The error bars represent the standard error of at least 3 replicates.
Table 3.1. Fast- and slow-relaxation components of NPQ (NPQF and NPQS, respectively) in two varieties of rice, At-354 and Bg-352 at the early vegetative and flowering stages subjected to 0, 50 and 100 mM NaCl. Measurements were carried out 14 days after initiating salt treatment. Leaves were exposed to actinic light of 2500 µmol m-2 s-1 for 20 minutes at 25 oC in the presence of 390 µL
-1 L CO2.
0 mM 50 mM 100 mM At-354 Bg-352 At-354 Bg-352 At-354 Bg-352 Early vegetative NPQF 1.17 ± 0.0057 1.15 ± 0.0046 1.4 ± 0.00172 1.38 ± 0.00673 1.48 ± 0.00243 0.55 ± 0.00653 NPQs 0.23 ± 0.0063 0.25 ± 0.0035 0.3 ± 0.00653 0.6 ± 0.00673 0.32 ± 0.00463 0.45 ± 0.00435
Flowering NPQF 1.55 ± 0.00674 1.53 ± 0.00854 1.58 ± 0.00654 1.4 ± 0.00545 2.02 ± 0.00865 1.85 ± 0.00743 NPQs 0.25 ± 0.00765 0.27 ± 0.0065 0.45 ± 0.00554 0.8 ± 0.00432 0.48 ± 0.00765 0.85 ± 0.00654
181 a b
d c
Figure 3.14. Chlorophyll fluorescence relaxation kinetics of control and salt treated rice plants at the flowering stage recorded using the PAM 101 fluorometer. (a) At-354 control plants and (b) AT-354 plants exposed to 100 mM NaCl. (c) Bg-352 control plants and (d) Bg-352 plants exposed to 100 mM NaCl. Measurements were carried out 14 days after initiating salt treatment. Leaves were exposed to actinic light of 2500 µmol m-2 s-1 for 20 minutes at 25 oC in the presence of 390 µL L-1
CO2. After that, the actinic light was switched off and a high intensity short duration light (1 second pulse of red light with a PFD of 4000 µmol m-2 s-1) was applied at 3 minutes intervals for over 45 minutes.
182 3.4.6. PSI Photochemistry
In addition to chlorophyll fluorescence, measurements of P700, the primary electron donor of PSI were performed using absorbance in the near infra red as an indicator of P700 redox state.
Measurements of the decay of P700 signal following a light-dark transition give information about
PSI electron transport. P700 became more oxidised when plants were exposed to salt (Figure 3.15 a, b). The redox state of P700 in control plants in both varieties is more reduced than the salt-treated plants at the early vegetative stage. In the salt-sensitive Bg-352 variety, P700 is more oxidised than in At-354 in both salt treatments. At the flowering stage, P700 in Bg-352 control plants is more oxidised than in At-354 control plants and it is similar to the oxidation of the At-354 plants treated with 50 mM NaCl. P700 in both varieties of plants became more oxidised at any given irradiance when exposed to higher salt concentrations.
The measure of the proportion of active PSI centres was also analysed. This measures PSI centres where P700 could be oxidised by light and then re-reduced in darkness relative to the total PSI oxidized by far red light. The At-354 control plants showed a higher proportion of active PSI centres than the Bg-352 control plants (Figure 3.15 c, d). Salt treated plants from both varieties showed higher proportion of active PSI centres than control plants. However, the proportion of active PSI centres in Bg-352 was higher than in At-354 salt-treated plants.
183 Early vegetative stage Flowering stage a b
d c
Figure 3.15. Redox state of P700 (a, b) and the proportion of 'active' PSI centres (c, d) of two rice varieties subjected to different salt concentrations plants at the early vegetative stage (a, c) and the flowering stage (b, d) of Bg-352 (closed symbols) and At-354 (open symbols) exposed to: 0 (black squares), 50 (blue triangles) and 100 (red diamonds) mM of NaCl. Leaves were exposed to actinic
o -1 light for 20 minutes at 25 C in the presence of 390 μL L CO2. The error bars represent the standard error of at least 3 replicates.
184 Measurements of the rate constant of P700 reduction gives information about the extent to which the electron transport chain is being regulated. Stepien and Johnson (2009) observed that the rate constant decreased in Arabidopsis thaliana and Thellungiella salsuginea when plants were subjected to salt. Consistent with these results, the rate constant decreased with increasing salt concentrations. At-354 and Bg-352 control plants at the early vegetative stage showed significantly higher rate constants than salt treated plants (Figure 3.16 a, b). At the flowering stage, the rate constant is notably higher in At-354 control plants than Bg-352 control plants.
The PSI electron transport rate (PSI ETR) of the flowering plants is higher than the early vegetative plants (Figure 3.16 c, d). PSI ETR in At-354 and Bg-352 plants increased in both stages when exposed to salt, in spite of the greatly inhibited rate of PSII electron transport. This is an indication of cyclic electron transport. However, at the flowering stage, Bg-352 treated with 100 mM showed a reduction in PSI ETR compared to the At-354.
185 Early vegetative stage Flowering stage a b
d c
Figure 3.16. The rate constant of P700 reduction (a, b) and PSI electron transport rate (PSI ETR) (c, d) plants at the early vegetative stage (a, c) and the flowering stage (b, d) of Bg-352 (closed symbols) and At-354 (open symbols) exposed to: 0 (black squares), 50 (blue triangles) and 100 (red diamonds) mM of NaCl. Leaves were exposed to actinic light for 20 minutes at 25 oC in the
-1 presence of 390 μL L CO2. The error bars represent the standard error of at least 3 replicates.
186 3.5. Discussion
Subjecting rice to a range of NaCl concentrations brought about a clear salinity responses in both
At-354 and Bg-352 varieties. In both At-354 and Bg-352, plant growth was decreased and leaf chlorosis occurred when exposed to salt (Figure 3.6 b, c). Apart from that, the leaf area showed a significant drop with increasing salt concentrations (Figure 3.7). These results emphasise that salinity causes a drop in biomass of rice plants. Effects of salt on plant growth occur due to both osmotic and ionic stress (Läuchli and Grattan, 2007; Munns and Tester, 2008). A reduction in plant growth due to osmotic stress is rapid and occurred within minutes of exposure to salt (Läuchli and
Grattan, 2007). Accumulation of excess Na+ in leaves over a period, caused toxic stress, which leads to an increase in leaf mortality with chlorosis and necrosis, and a decrease in photosynthesis
(Yeo and Flowers, 1986; Glenn et al., 1999). This caused a reduction in overall plant growth
(Munns, 2002; Läuchli and Grattan, 2007). Decreases in the leaf area of rice under salinity were also observed in a study performed by Hakim et al. (2014a). The early vegetative stage or the tillering stage is the initial stage of leaf and tiller development (Zeng and Shannon, 2000).
Reduction in the tiller formation of rice at the early vegetative growth stage affects the final yield
(Hoshikawa, 1989).
Rice plants at the flowering stage also showed a significant decreases in leaf area when exposed to salt, however, it was not as prominent as at the early vegetative stage. This indicated that plants at the flowering stage showed more tolerance to salt. Several studies have shown that majority of the annual crops are tolerant to salinity at the germination stage but are sensitive during the emergence and the early vegetative stage (Läuchli and Epstein, 1990; Maas and Grattan, 1999; Läuchli and
Grattan, 2007). Studies by Flowers and Yeo (1981) and Lutts et al. (1995) showed that, rice is more tolerant at the reproductive stage than the young seedling stages. In contrast, a study by Asch et al.
187 (2000) showed that, from all growth stages, the most sensitive to salinity is the panicle initiation of the reproductive stage. Another study found that rice is less sensitive to salt at germination, tillering and grain filling stages, but sensitive at both seedling and reproductive stages (Lafitte et al., 2004).
Khatun et al. (1995) showed that salt stress delayed flowering, reduced the number of productive tillers, the number of fertile florets per panicle, the weight per grain and the grain yield, therefore, effects on grain yield were very much more severe than on vegetative growth. According to the previous studies and current results, it is evident that the sensitivity to salinity at different growth stages depends on the rice variety. Therefore, improvement of salt tolerance of different rice varieties should target the specific growth stages that are more sensitive to salt which can substantially affect grain yield (Walia et al., 2005).
Leaf chlorophyll content can be used as an indicator of leaf injury in stressed plants (James et al.,
2002). Decreases in leaf chlorophyll content under salinity have been observed in various crops, including sunflower (Ashraf and Sultana, 2000; Akram and Ashraf, 2011), alfalfa (Winicov and
Seemann, 1990), wheat (Arfan et al., 2007; Perveen et al., 2010) and castor bean (Pinheiro et al.,
2008). Salt had a significant impact on leaf chlorophyll content in the salt-sensitive Bg-352 at both stages (Figure 3.8 a, b). Loss of chlorophyll in At-354 at the early vegetative stage was not prominent at low salt concentrations. However, at the flowering stage, both varieties showed a gradual decrease in the total chlorophyll content. Previous studies have shown a decrease in leaf chlorophyll content in rice under salinity (Lutts et al., 1996; Hakim et al., 2014b; Senguttuvel et al.,
2014). In these studies, salt sensitive rice varieties showed marked reduction in chlorophyll content compared to salt tolerant varieties. The possible reasons for the decline in chlorophyll content in salt stressed plants could be the activation of chlorophyllase enzymes or the disruption of the structure of pigments and protein complexes by accumulation of sodium and chloride ions (Reddy and Vora,
1985; Fang et al., 1998; Djanaguiraman and Ramadass, 2004).
188 The measure of the chlorophyll a/b ratio indicates changes in the composition of the thylakoid membrane and positively correlates with the ratio of reaction centre cores to light harvesting chlorophyll-protein complex (LHC) (Terashima and Hikosaka, 1995). A study by Djanaguiraman et al. (2006) showed that the degradation of chlorophyll-b is more severe than chlorophyll-a, which results in an increase in chlorophyll a/b ratio when plants are exposed to salt. LHCII contains the majority of chlorophyll-b and has a low chlorophyll a/b ratio (1.3-1.4) compared to chlorophyll binding proteins associated with PSII core which binds no chlorophyll-b (Evans, 1989; Green and
Durnford, 1996). Therefore, increases in chlorophyll a/b indicates either loss of light harvesting complexes (LHCs) relative to the reaction centres (RC) in photosystems, or loss of PSII compared to PSI or a combination of both (Anderson, 1986). The increase in the chlorophyll a/b ratio in At-
354 treated with 50 mM NaCl indicates that salt decreases the light harvesting complexes compared to the reaction centres possibly in line with a change in PSI: PSII ratio or in relative antenna size.
Decreases in the chlorophyll a/b ratio at high salt concentrations indicate that salt affects the reaction centres (or preferentially PSI) in both varieties, resulting in a decrease in photosynthesis.
Similar results were observed in studies performed by Lutts et al. (1996) and Senguttuvel et al.
(2014). The increase in chlorophyll a/b possibly reflects a controlled breakdown of LHC complexes, whilst a decrease is probably an indication of a less controlled, stress induced, loss of reaction centres. This could be examined further using a biochemical analysis of thylakoid composition
Gas exchange parameters, including CO2 assimilation rate (A) stomatal conductance (gs) and transpiration rate (T) were significantly decreased in both salt tolerant At-354 and salt sensitive Bg-
352 when exposed to 50 and 100 mM of NaCl (Figure 3.9). The same effect was observed in the salt tolerant Pokkali and moderately salt tolerant IR-2153 varieties when exposed to salt
189 concentrations higher than 100 mM of NaCl (Yeo et al., 1991). Another study showed that, adverse effects of salt on photosynthetic rate was associated with with a significant decrease in the stomatal conductance in all rice varieties (Hakim et al., 2014a). Sengutthuvel et al. (2014) showed a decrease in stomatal conductance and transpiration rate in both salt tolerant and sensitive cultivars under salinity stress. However, salt tolerant genotypes showed higher stomatal conductance compared to the salt sensitive genotypes. Decreases in stomatal conductance are among the most responsive salinity tolerance mechanisms in rice (Moradi and Ismail, 2007). Under control conditions, CO2 assimilation and stomatal conductance were higher in the salt-tolerant At-354 than in Bg-352. This implies that salt tolerance in At-354 is not achieved by preventing the flow of salt into the plant via the lowering transpiration stream. Rather, leaf osmotic potential is maintained below the soil water potential and salt is either prevented from entering the cells or tolerated more than in the sensitive
Bg-352 variety. However, at the flowering stage At-354 control plants showed a higher stomatal conductance compared to the vegetative stage.
Exposure to salt, resulted in stomatal closure, which causes low conductance (Farquhar et al.,
1982a; Farquhar et al., 1982b; Downton et al., 1985). This decline is mirrored in the reduction of assimilation in both rice varieties. Excess salt affects photosynthesis in two ways: (1) low rate of
CO2 diffusion (flux) of into leaf, caused by reductions in stomatal and mesophyll conductance
(stomatal limitations) and (2) disruption of metabolic potential for photosynthesis (nonstomatal limitations) (Chaves, 1991; Tezara et al., 1999; Ashraf, 2003; Lawlor and Tezara, 2009; Chaves et al., 2011). Stomatal limitations are considered as the major contributor which inhibits photosynthesis (Cornic and Briantais, 1991). Dionisio-Sese and Tobita (2000) observed a reduction in carbon assimilation rate and stomatal conductance in rice due to stomatal closure rather than nonstomatal inhibition of photosynthesis. However, studies have shown that nonstomatal limitations occurred through direct effect of NaCl on photosynthetic apparatus and responsible for low
190 photosynthetic rates in plants (Ball and Farquhar, 1984; Seeman and Critchley, 1985; Seeman and
Sharkey, 1986; Tezara et al., 2002; Chaves et al., 2009). Therefore, response curves between CO2 assimilation (A) and internal CO2 concentration (Ci) were plotted to differentiate limiting effects on photosynthesis into stomatal and nonstomatal factors (Farquhar and Sharkey, 1982). According to the results from the ACi curves (Figure 3.10), the decrease in assimilation in salt tolerant At-354 is due to lack of CO2 inside leaves (closure of stomata) rather than damaging the components of the thylakoid membranes. In Bg-352, the drop in the rate of assimilation cannot restored by increasing the external CO2 concentration. This inhibition of CO2 assimilation has been attributed to a reduced
Rubisco activity, RuBP regeneration and triose phosphate utilization or to increased sensitivity of
PSII to NaCl (Ball and Farquhar, 1984; Ball and Anderson, 1986; Seeman and Sharkey, 1986;
Johnson and Murchie, 2011). Measurements of foliar NaCl in salt-treated rice plants would give a clearer idea of whether the non stomatal limitation was due to salt accumulation in Bg-352 is more than in At-354. Moradi and Ismail (2007) found that, two tolerant rice breeding lines have lower
Na+ concentration and higher K+/Na+ ratio in leaves than the salt sensitive cultivar during both seedling and reproductive stages. Dionisio-Sese and Tobita (2000) observed a significant increased in Na+/K+ ratio in salt sensitive rice cultivars with increasing salt concentrations than salt tolerant
Pokkali. Differential distribution of Na+, Cl- and K+ was previously observed in rice leaves (Yeo and Flowers, 1982; 1984; Yeo et al., 1985; Aslam, 1987) and concentrations were much higher in old leaves than young leaves (Wang et al, 2012). Sengutthuvel et al. (2014) found that salt sensitive rice genotypes reduced photosynthetic carbon assimilation due to both stomatal and nonstomatal limitations.
The maximum quantum efficiency of PSII, Fv/Fm is the most extensively measured parameter which indicates the tolerance or sensitivity of a plant to stress (Peñuelas and Boada, 2003; Siddiqui et al., 2014). Studies by Dionisio-Sese and Tobita (2000) and Moradi and Ismail (2007) found that,
191 Fv/Fm was not affected by salt in either salt tolerant or salt sensitive rice cultivars. However, salt stress significantly reduced the efficiency of photosynthesis in both At-354 and Bg-352 rice genotypes during the early vegetative and the flowering stages (Figure 3.11). Decrease in Fv/Fm is a clear indication that PSII was affected by salt and photoinhibition was occurring. However, the decrease in Fv/Fm was smaller in At-354 than in Bg-352 at both growth stages. This effect was also observed in a study performed by Senguttuvel et al. (2014), where salt sensitive cultivar, IR29 had a lower Fv/Fm ratio when exposed to 120 mM NaCl, compared to the salt tolerant IR72593. Also, the efficiency of PSII (ΦPSII) and the rate of electron transport (PSII ETR) in At-354 was higher than in Bg-352, even when plants were stressed (Figure 3.12). With this we can conclude that the Bg-352 variety is more sensitive to salinity than the At-354 variety. This is reflected in photosynthesis of
Bg-352 being inhibited at lower salt concentrations compared to At-354. Similar results were observed in a study performed by Moradi and Ismail (2007) and García Morales et al. (2012). They showed that, there was no reduction in ΦPSII and PSII ETR in the tolerant lines, but salt-sensitive cultivars showed a significant decrease in ΦPSII and PSII ETR under salt stress. It is noteworthy that, in this study, the increase in chlorophyll a/b in salt tolerant AT-354 at 50 mM is consistent with the down-regulation electron transport in PSII indicated by both Fv/Fm (Figure 3.11) and ΦPSII
(Figure 3.12) compared to salt sensitive Bg-352. However, the decrease in chlorophyll a/b ratio in both rice varieties at high salt concentration suggests a preferential loss of reaction centres, possibly reflecting damage to these. This should be examined using biochemical analyses of thylakoid membranes.
Plants adapt regulatory processes to overcome the excitation pressure in electron transport under abiotic stresses and protect plants from toxic ROS. Non-photochemical quenching (NPQ) is a photoprotective mechanism in photosynthesis which protects the components of PSII by dissipating excess energy as heat when plants were exposed to stress (Niyogi et al., 1998; Müller et al., 2001;
192 Rohácek, 2002; Golding and Johnson, 2003; Netondo et al., 2004; Redondo-Gómez et al., 2006;
Tezara et al., 2008; Li et al., 2008; Ribeiro et al., 2009; Stepien and Johnson, 2009; Silva et al.,
2011). Moradi and Ismail (2007) showed an increased NPQ in both salt tolerant and salt sensitive rice cultivars. Another study showed that, NPQ increased rapidly with an increase in light intensity and no further significant increase in NPQ was observed above 700 µmol photons m-2 s-1 in both salt sensitive rice cultivar and salt tolerant cabbage cultivar (Zhu et al., 2011). Consistent with that,
NPQ is relatively higher in the salt-treated AT-354 and Bg-352 plants than in the control plants.
However, at the early vegetative stage, Bg-352 plants treated with 100mM salt showed a lower
NPQ than the control plants and this was not observed at the flowering stage. The reason for this drop might be that at early stages, Bg-352 plants have lower ability to generate a pH gradient across the thylakoid membrane when plants are under severe stress conditions. The drop in the NPQ in Bg-
352 at high salt concentration suggested a possible uncoupling of the thylakoid membranes induced by the high levels of ions. Measurements of intracellular ion concentrations and measurements on isolated membranes would allow this to be tested.
Chlorophyll fluorescence level (Fm' in Figure 3.2) of the steady-state photosynthesis is always below the maximum fluorescence level (Fm in Figure 3.2) due to both photochemical and non- photochemical quenching processes (Roháček et al., 2008). The non-photochemical processes include the generation of a pH gradient gradient across the thylakoid membranes, inactivation and/or photodamage of PSII reaction centres, state transitions, zeaxanthin formation through the xanthophyll cycle activation and conformational changes within thylakoid membranes (Bilger and
Björkman, 1990; Ruban and Horton, 1995; Govindjee, 1995; Pospíšil, 1998; Maxwell and Johnson,
2000). After switching off the actinic light, the relaxation of nonphotochemical process occurred
(Roháček et al., 2008). The pH dependent photoprotective, high energy state quenching (qE) relaxes within seconds to minutes, state transitions (qT) relaxes within tens of minutes and photoinhibition
193 (qI) relaxes over a long period of time (hours) (Walters and Horton, 1991; Maxwell and Johnson,
2000). However, qE and qT cannot easily be distinguished from their relaxation kinetics. Therefore, relaxation analysis were performed to distinguished, qE and qI components of NPQ. According to results (Table 3.1), in both rice varieties, the majority of quenching relaxed rapidly in the dark, showing that it was high energy-state quenching (qE) rather than photoinhibition (qI). However, compared to the salt-treated At-354, increased in slow relaxing component in salt-treated Bg-352 suggested the occurrence of qI at high salt concentration. Stepien and Johnson (2009) observed a similar effect in salt sensitive Arabidopsis thaliana compared to salt tolerant Thellungiella salsuginea when exposed to 150 mM of NaCl.
PSI photochemistry was analysed by measuring the kinetics of re-reduction of P700+ following a light to dark transition (Clarke and Johnson, 2001). According to results of the redox state and the turnover of P700, it is evident that P700 become more oxidized in both rice varieties when exposed to salt (Figure 3.15 a, b). However, P700 became more oxidized in Bg-352 than in At-354 at both growth stages. Similar results were observed in salt sensitive Arabidopsis thaliana when exposed to salt (Stepien and Johnson, 2009). PSI was also analysed by measuring the relative concentration of active PSI centres which of can be oxidized by light and then re-reduced in the dark. The proportion of 'active' PSI showed a marked increase in both varieties when exposed to salt. According to a study by Klughammer and Schreiber (1994), PSI should be more inactive when the acceptor side is reduced under abiotic stress conditions. The increase in the 'active' PSI centres under stress conditions was previously observed in the studies carried out by Golding and Johnson (2003) and
Stepien and Johnson (2009) on barley under drought stress. These studies suggested that the
'additional' PSI centres are involved in the cyclic electron transport which generates a pH gradient to maintain NPQ. Salt-treated Bg-352 plants maintained a higher relative concentration of active
PSI than the control plants in both stages, which suggests that cyclic electron transport might be
194 more prominent in Bg-352 than At-354 under salt stress. However, Bg-352 showed a substantial drop in NPQ under high salt concentration at the early vegetative stage. The possible reason for the marked drop in NPQ in the salt-sensitive rice variety might be due to leakage of proton through membranes damaged by the high levels of salt ions.
PSI turnover can be examined by measuring the re-reduction of P700+. Fitting P700+ decay curves with a single exponential decay curve yields a pseudo first order rate constant (k) (Harbinson and
Woodward, 1987). The rate constant of PSI electron transport gives information about the extent to which the electron transport chain is being regulated (Stepien and Johnson, 2009). The rate constant values decreased with increasing salt concentrations in both rice varieties. However, the decrease is more substantial in Bg-352 than in At-354. The product of the rate constant and oxidized P700+ gives the rate of electron transport to PSI (Clarke and Johnson, 2001; Golding and Johnson, 2003).
Despite there being a marked reduction in PSII ETR, the highest electron transport rate of PSI (PSI
ETR) was observed in stressed plants in both stages. However, at the flowering stage, PSI ETR in salt-stressed Bg-352 variety decreased. This might be due to the drop of PSI centres under salinity.
At the early vegetative stage, salt-treated Bg-352 plants showed a higher PSI ETR than At-354.
However, at the flowering stage, PSI ETR of salt-treated At-354 plants was higher than in Bg-352.
Plants absorb inorganic chemicals from the soil solution most of which are essential for plant growth, but some, like Na+ and Cl- are non-essential or toxic if absorbed in excess (Rengasamy,
2010). However, at relatively low concentrations, plants can restrict the excessive ion uptake and all plants control Na+ and Cl- to some extent. Plants exhibit several mechanisms to survive in saline environmental conditions such as, salt exclusion at the roots, salt transport prevention to the leaves, salt elimination by leaf shedding and salt excretion at the leaves (Lambert and Turner, 2000). A study by Stepien and Johnson (2009) showed that halophyte Thellungiella salsuginea has relatively
195 low accumulation of Na+ and maintains Na+/K+ ratio compared to Arabidopsis thaliana. Another study showed that, Mesembryanthemum crystallinum (common ice plant), which is a halophyte, has epidermal bladder cells to maintain Na+/K+ in the leaves (Agarie et al., 2007). Yeo and Flowers
(1986) showed that, although 99% of arriving Na+ was sequestered into the expanded rice leaves during salt stress, the apoplastic Na+ concentration could reach 500 mM within 7 days resulting cell dehydration and stomatal closure. This was also observed in the salt tolerant rice variety, Pokkali
(Krishnamurthy et al., 2009; Krishnamurthy et al., 2011). In this study, we observed the Bg-352 is more salt sensitive than At-354. This might be due to high levels of salt ions accumulation in Bg-
352 than At-352. Therefore, it is important to examine the intracellular ion concentrations in salt treated leaves in both rice varieties. To fully elucidate the changes in the ratio of Na+/K+ in At-354 and Bg-352, further analysis needs to be done.
In summary, our physiological analysis of photosynthesis indicates that the regulation of photosynthesis in the salt-tolerant At-354 is more prominent than the salt-sensitive Bg-352 when plants were exposed to salt. Exposure of Bg-352 to salt resulted in substantial decreases in gas exchange, PSII photochemistry and loss of chlorophyll. The decrease in photosynthesis in AT-354 is caused by stomatal limitations which restrict CO2 entry into the plants whereas the decrease of photosynthesis in Bg-352 is caused by non-stomatal limitations such as damage to membranes and proteins. Results suggested that At-354 protects PSII centres from excitations pressure by down- regulating the electron transport chain and maintaining a pH gradient by cyclic electron transport associated with PSI to support NPQ. At high salt concentration, this regulation starts to fail in Bg-
352. The failure might result from a specific loss of PSI, resulting in reduced cyclic electron flow or an increase in the leakiness of the thylakoid membranes resulting in loss of pH gradient. In the future, thylakoids should isolated from the salt-treated rice plants to measure the leakiness of the membranes and to quantify PSI and PSII.
196 Chapter 4
Regulation of Photosynthesis in Thellungiella salsuginea under abiotic stress
Sashila Abeykoon Walawwe
Giles N. Johnson
197 Preface
Sashila Abeykoon Walawwe is the primary author of this paper.
Plant growth by Sashila Abeykoon Walawwe
Photosynthetic measurements of the activity of PTOX by Sashila Abeykoon Walawwe
SDS-PAGE, BN-PAGE and immunoblot analysis by Sashila Abeykoon Walawwe rt-PCR analysis by Sashila Abeykoon Walawwe
Mass spectroscopy by the Protein Mass Spectroscopy core facility centre in the University of
Manchester.
198 4.1. Abstract
Thellungiella salsuginea is an extremophile able to grow and reproduce in extreme environments.
Stepien and Johnson (2009) identified a protein, known as plastid terminal oxidase (PTOX), which acts as an alternative electron sink in this plant under salt stress. When plants are stressed with high salt concentrations, PTOX diverts electrons from plastoquinol to oxygen. T. salsuginea plants were cultivated on soil and challenged with abiotic stresses other than salt, specially drought, different growth irradiances, cold and cold combined with high light for 14 days. In addition, plants were also grown under semi-natural conditions in a greenhouse. T. salsuginea leaves exposed to abiotic stress conditions were tested for PTOX protein content and upregulation of PTOX gene transcripts under salinity stress were compared to the control plants. Efficiency of PSII (ΦPSII) and the relative electron transport of PSII (PSII ETR) were also measured under 2% and 20% O2 concentrations.
Direct electron transport from PSII to PTOX and then to oxygen via the PQ pool accounted for up to 30% of total PSII electron flow in T. salsuginea (Stepien and Johnson, 2009). Efficient electron flow from PSII to PTOX would however, probably require co-location of these complexes in the same thylakoid fraction. To examine the location of PTOX in the thylakoid membrane, immunoblot analyses were performed, to test for changes in other protein complexes which may be associated with PTOX. In addition blue-native polyacrylamide gel electrophoresis and immunoblots were performed to isolate and detect the PTOX protein with any associated complexes. Increases in relative PTOX protein abundance, upregulation of PTOX gene transcripts and activity of PTOX under abiotic stresses, including salt, drought, cold combined with high light, different growth irradiances and plants grown in a greenhouse indicated the involvement of PTOX in stress regulation in T. salsuginea. However, this was not observed in plants treated with cold alone.
Although immunoblot analysis showed a prominent signal, mass spectrometry data did not allow identification of PTOX. This results suggests that further studies are needed to identify the precise localisation of the PTOX protein in the thylakoid membranes in T. salsuginea.
199 4.2. Introduction
It is essential that the energy and reducing power produced by photosynthetic electron transport be kept in balance with the requirements of CO2 fixation to prevent reactive oxygen species (ROS) production (Trebst et al., 2002; Triantaphylidés et al., 2008; Triantaphylidés and Havaux, 2009).
Abiotic stresses, including salinity, drought, cold and high light, can cause an energy imbalance between electron transport and CO2 fixation in plants, which can trigger oxidative stress and encourage the production of ROS such as superoxides and singlet excited oxygen (Aro et al., 1993;
Foyer et al., 1994; Asada, 2000; Vass and Cser, 2009; Vass, 2012). ROS are highly active and damage many cellular components, including membranes, proteins and DNA. Although plants produce various enzymes and antioxidants to scavenge ROS and protect themselves from oxidative damage, production of these is energetically demanding (Asada, 2000). Therefore, plants regulate the electron transport chain (ETC) of photosynthesis as an alternative strategy which is energetically less demanding and inhibits the production of ROS (Ott et al., 1999; Golding and
Johnson, 2003; Stepien and Johnson, 2009).
In addition to having photoprotection processes, including non photochemical quenching, alternative pathways of electron transport play a vital role in regulation of electron transport when plants are challenged with environmental stresses (Johnson, 2005). Cyclic electron transport (CET) around PSI in the thylakoid membranes is the best known alternative electron transport, associated with the regulation of the electron transport pathway. Early studies have shown that CET is involved in electron transport in C3 plants only under stress conditions (Herbert et al., 1990).
However, it has been found that CET is an essential pathway involved in a wide variety of photosynthetic organisms, including higher plants, cyanobacteria and algae, which supports growth and development (Herbert et al., 1990; Fork and Herbert, 1993; Bendall and Manasse, 1995; Heber,
2002; Allen, 2003; Munekage et al., 2004).
200 In addition to CET, the Mehler reaction, or the water-water cycle, occurs at the acceptor side of PSI and directs excess electrons from PSI to oxygen when PSI acceptors are depleted (Asada, 1999;
Clarke and Johnson, 2001; McDonald et al., 2011). Electrons accepted by oxygen produce ROS which disproportionates to H2O2 and O2, catalyzed by superoxide dismutase (SOD). H2O2 is further reduced to water by ascorbate peroxidase (Asada, 1999). Although the Mehler reaction has been suggested to act as a safety process, to divert excess electrons from the electron transport, it has several drawbacks which probably outweigh any potential benefits. Reduction of molecular oxygen, forms harmful radical species which can damage PSI and PSII (Clarke and Johnson, 2001). Apart from that, producing scavenging enzymes in higher concentrations is energetically demanding for plants. According to several studies, the Mehler reaction is insufficient to provide significant protection from photoinhibition (Cornic and Briantais, 1991; Wiese et al., 1998; Clarke and
Johnson, 2001).
Chlororespiration is another electron transfer pathway in which stromal reducing equivalents are transferred to dioxygen through the PQ pool (Bennoun, 1982). In this pathway, the transfer of electrons from plastoquinol to oxygen is mediated by a chloroplast targeted terminal plastoquinol oxidase known as the plastid terminal oxidase (PTOX). NAD(P)H dehydrogenase (NDH) and plastid terminal oxidase (PTOX) are the major components involved in this non-photochemical reduction of the PQ pool (Carol et al., 1999; Wu et al., 1999; Rumeau et al., 2007). NDH complex, which is involved in both CET and chlororespiration, consists of large number of chloroplast and nuclear-encoded subunits (Rumeau et al., 2007). Mitorespiration and photosynthesis interact with each other through ATP synthesis, reducing potential and metabolite exchange (Hoefnagel et al.,
1998). Due to this, it was difficult to differentiate the activity of mitochondrial oxidases and chloroplast oxidases involved in the PQ oxidation (Bennoun, 1994; Bennoun, 1998; Bennoun,
2001). The identification of propyl gallate as a potent inhibitor of the chlororespiratory oxidase,
201 provided important information on chlororespiration and the activity of PTOX as the ultimate component of this pathway which mediates the transfer of electrons from plastoquinol to oxygen
(Cournac et al., 2000a; Cournac et al., 2000b; Josse et al., 2000).
PTOX is a plastid localized plastoquinol oxidoreductase, commonly found in photosynthetic organisms, including green algae and higher plants (Carol and Kuntz, 2001; Kuntz, 2004; Sun and
Wen, 2011). Studies have shown that PTOX is encoded by a single gene in higher plants and by two genes in some eukaryotic algae (Wang et al., 2009). PTOX is proposed to be an interfacial membrane protein with a di-iron carboxylate centre in the active site where iron is the catalytic co- factor (Kuntz, 2004). PTOX shows sequence similarity to plant alternative oxidase (AOX) involved in an alternative pathway in mitochondrial respiration which mediates electron transfer from ubiquinol to oxygen (Kuntz, 2004; McDonald et al., 2011; Sun and Wen, 2011). PTOX was first identified in the Arabidopsis thaliana (A. thaliana) pigment mutant, immutans, and the tomato ghost mutant. Both these mutants, which are impaired in the PTOX gene, show variegated phenotypes, with clear white and green sectors under low-moderate light while, under high light conditions, leaves bleach due to photo-oxidative damage (Kuntz, 2004; McDonald et al., 2011; Sun and Wen, 2011). Further studies have shown that bleached leaves in both immutans and ghost contained low carotenoid content and accumulate phytoene in leaves, which suggests that PTOX is involved in carotenoid biosysnthesis. PTOX is known to be involved in the phytoene desaturation in carotenoid biosynthesis pathway. Therefore, lack of PTOX will lead to the blockage in carotenoid production in plants (Sun and Wen, 2011).
An induced level of PTOX, in plants challenged with different abiotic stresses, was observed in several species (Rumeau et al., 2007). A study on oat (Avena sativa) showed high level of PTOX and NDH under high light and heat treatment (Quiles, 2006). Lodgepole pine also showed an
202 elevated level of PTOX during acclimation to winter cold (Savitch et al., 2010). A study by Kong and co-workers (2003) showed that, a salt tolerant rice variety showed an increased level of PTOX and two splicing mechanisms in PTOX gene expression. They found that, OsIM1 transcript showed
66% amino acid sequence similarity to tomato PTOX and was increased by salt and abscisic acid treatment. Brassica fruticosa showed high levels of stress tolerance and more elevated levels of
PTOX than Brassica oleracea when exposed to high light and heat (Díaz et al., 2007). Several studies have shown that plants exhibiting weak chloroplast antioxidant systems showed an increased PTOX level when exposed to stress. For example, studies have detected an induced level of PTOX in high mountain plant Rananculus glacialis which has low antioxidative scavenging capacity and low NPQ (Streb et al., 2005; Laureau et al., 2013). Another study by Rizhsky and co- workers (2002) showed double antisense tobacco plants lacking two major hydrogen peroxide detoxifying enzymes, catalase and ascorbate peroxidase, showed elevated levels of PTOX. These studies suggested that PTOX act as an electron safety valve which prevents over reduction of the
PQ pool under high light.
In addition, the activity of PTOX under various stress conditions were observed in other photosynthetic organisms. It was found that PTOX gene is present in all high light adapted ecotypes of Prochlorococcus marinus but not in the cyanobacterial strains found in low light and low temperature environment (Rocap et al., 2003; Kettler et al., 2007; Luo et al., 2008). A study has shown that Synechococcus WH8102, a marine cyanobacterium possess an alternative electron flow to O2 via PTOX when PSI activity is limited due to low iron levels. They hypothesized that
Synechococcus uses PTOX, which only has two iron atoms rather cytochrome b6f and PSI which have 18 iron atoms altogether to survive in low iron conditions (Bailey et al., 2008). Apart from that, expressed sequence tags data suggested that PTOX was transcribed in two diatoms
Phaeodactylum tricornutum and Thalassiosira pseudonana under low iron conditions (McDonald et
203 al., 2011). Studies on the green alga, Haematococcus pluvialis showed that changes in the PTOX transcripts occurred under various stresses, including high light, excess iron or salt and low temperature (Wang et al., 2009; Li et al., 2008; Li et al., 2010). These studies also showed that
PTOX is involved in the production of astaxanthin and plays a protective role against stress. Work from Cardol et al. (2008) showed a deep sea/low light strain of the green picoeukaryote
Ostreococcus strain (RCC 809), which lives in low iron conditions, lacks PSI compared to surface/ highlight strain (OTH95) bypasses electrons in a water-water cycle to generate a pH gradient across the thylakoid membranes. This cycle passes large numbers of electrons through PSII to oxygen with the involvement of PTOX (Cardol et al., 2008). Increased levels of PTOX transcripts in phosphorus starved cells of Chlamydomonas reinhardtii suggested that, PTOX plays a major role in stress responses in this photosynthetic algae (Moseley et al., 2006). Apart from the photosynthetic organisms, cyanopages, viruses which infects cyanobacteria, possess PTOX genes in their genomes.
For example, cyanophage Syn9 consists of photosynthetic genes including, plastocyanin, PTOX,
PsbA, PsbD (Weigele et al., 2007). It has been hypothesized that these genes provide a photoprotective function during the phage propagation (Lindell et al., 2005; Weigele et al., 2007).
A study by Stepien and Johnson (2009) detected induced PTOX levels in Thellungiella salsuginea plants under salt stress. They also observed that PTOX acts as an alternative electron sink, where transfer of excess electrons from over-reduced plastoquinol pool to oxygen provides a protection from ROS. They concluded that in salt-stressed Thellungiella salsuginea, electron transport to plastid terminal oxidase accounted for up to 30% of total PSII electron flow. Thellungiella salsuginea (T. salsuginea) is an extremophile, which can grow and reproduce in many adverse weather conditions. Therefore, we can assume that the alternative pathway mediated by PTOX might support the regulation of electron transport chain under abiotic stresses other than salinity.
However, it has not been shown whether PTOX is expressed when plants are exposed to other
204 stresses. Therefore, this study is focused on examining the expression and the activity of PTOX when T. salsuginea plants are challenged with drought, cold, different growth irradiances, cold combined with high light and plants grown in semi natural conditions.
PTOX level in leaves can change in response to changes to various abiotic stresses. This can be observed by examining the transcriptional regulation of genes. Therefore, this study also focuses on detecting the regulation of the PTOX gene transcript under salt stress. Lennon et al. (2003) found that, PTOX is bound to thylakoids, localised mostly in stromal lamellae with only a small amount found in the grana in spinach. Efficient electron flow from PSII to PTOX would however, probably require co-location in the same thylakoid fraction. Therefore, the precise location and the orientation relative to the plastoquinol pool is essential to understand the role of PTOX in photoprotection of stressed plants. Heyno and co-workers (2009) showed that PTOX encourages photo-oxidative stress when over-expressed in tobacco plants rather than photoprotection. Another similar study by Ahmad et al. (2012) showed that over-expression of algal PTOX in tobacco chloroplasts make plants more sensitive to high light than the wild type. Failure to induce the alternative pathway in these transgenic plants suggests that PTOX may be a subunit of some larger thylakoid protein complex. Therefore, this study also focused on detecting the effect of abiotic stresses on other complexes including PSII and cytochrome b6f and on attempting to isolate PTOX as a native protein along with any complexes associated with it.
205 4.3. Materials and Methods
4.3.1. Plant growth
Seeds of T. salsuginea (Thellungiella salsuginea; ecotype Shandong wild type) were stratified at 4 oC for three days and then germinated at a light intensity of 140 µmol m-2 s-1 provided from white fluorescent bulbs, day time temperature was 20 o C and night time temperature was 15 oC. 2-week old seedlings were transferred to pots filled with peat-based compost (except for plants used for drought treatment; used John Innes No 1 soil-based compost). 7-week old plants were used for the treatments.
4.3.2. Treatments
1) Drought
Plants were grown in a controlled environment growth room with a photon flux density of 140
µmol m-2 s-1 (Light Meter, SKYE Instruments LTD, UK) provided by white fluorescent bulbs on a
16 hour light/ 8 hour dark cycle. The day time temperature was 20 oC and night time temperature was 15 oC. 7-week old plants were supplied with 50 ml of water (for each pot) on the first day and measurements were taken after 14 days of the treatment.
2) Effects of different growth irradiances
Plants were grown from the seedling stage in a controlled environment growth room with a light intensity of 100 µmol m-2 s-1 (low light) provided by white fluorescent bulbs on 8 hour light/ 16 hour
206 dark cycle. The day time temperature was 20 oC and night time temperature was 15 oC. 7-week old plants were transferred to 400 µmol m-2 s-1 (moderate light- different shelf in the same growth cabinet fitted with compact fluorescent tubes). After 14 days, measurements were taken.
3) Cold
Plants were grown in a controlled environment growth room with a photon flux density of 140 µmol m-2 s-1 provided by white fluorescent bulbs on a 16 hour light/ 8 hour dark cycle. The day time temperature was 20 oC and night time temperature was 15 oC. 7-week old plants were then transferred to a growth room with a light intensity of 140 µmol m-2 s-1 and temperature of 4 oC. After
14 days measurements were taken.
4) Cold and high light
Plants were grown in controlled environment growth room with a photon flux density of 140 µmol m-2 s-1 provided by white fluorescent bulbs on a 16 hour light/8 hour dark cycle. The day time temperature was 20 oC and night time temperature was 15 oC. 7-week old plants were transferred to a growth room with 4 oC and light intensity of 1000 µmol m-2 s-1 (high light) provided by LED lamp.
After 14 days measurements were taken.
5) Salinity
Plants were grown in controlled environment growth room with a photon flux density of 140 µmol m-2 s-1 provided by white fluorescent bulbs on a 16 hour light/8 hour dark cycle. The day time temperature was 20 o C and night time temperature was 15 o C. 7-week old plants were treated with
207 250 mM of NaCl 14 days before taking measurements.
6) Plants grown in semi -natural green house
Seeds of T. salsuginea were stratified at 4 oC for three days and then transferred to an unheated greenhouse without supplementary lighting during the periods March-June 2011 and June-
September 2011 at botanical grounds in the University of Manchester. 2-week old plant seedling were transferred to pots filled with John Innes No 1 soil-based compost. 7-week old plants were used for the measurements.
4.3.3. Measuring chlorophyll fluorescence and P700 with 2% and 20% O2 concentrations
2% and 20% of O2 gas were supplied by mixing compressed oxygen and nitrogen gases from cylinders (BOC Gases) and using an MKS Mass Flow controller (MKS Instruments Inc.) to control the gas outlet (Clarke and Johnson, 2001; Stepien and Johnson, 2009). Thellungiella leaves were exposed to 2% and 20% O2. The activity of the plastid terminal oxidase was analysed by comparing the efficiency of PSII (ΦPSII) and the electron transport rate of PSII (PSII ETR) (described in
Chapter 2, Section 2.3.2.2) at the different O2 concentrations at different light intensities provided by a Luxeon III red LED in a laboratory built lamp and light intensity was measured using a PAR meter (SKP215; Skye Instruments, Powys, UK). An atmosphere of saturating CO2 was obtained by
-1 bubbling gas through a solution of 1 M Na2CO3/NaHCO3 (pH 9) (>1200 μL L CO2) (as described in Clarke and Johnson, 2001).
208 4.3.4. Isolating thylakoids from T. salsuginea leaves
Leaves of T. salsuginea (7 g) were collected, washed and blotted dry. The glass bowl of a Magimix food blender and glassware were pre-cooled in a freezer. Ice-cold grinding medium (25 ml), containing sorbitol (330 mM), HEPES (20 mM), NaCl (10 mM) and MgCl2 (5 mM) (pH 7.6,
NaOH) was partly thawed and leaves were ground briefly using the blender in slushy grinding medium. The solution was filtered through two layers of muslin and then through two layers of muslin and one layer absorbent cotton wool. The filtrate was centrifuged at 3000 xg for 5 minutes (5 o C). The pellet was washed with washing medium made from HEPES (5 mM) and MgCl2 (5 mM)
(pH 7.6) and re-centrifuged at 3000 xg for 5 minutes. The pellet was then resuspended in 1 ml of grinding medium, flash frozen and stored in the freezer until use.
4.3.5. Extraction of proteins from the thylakoid suspension
100 µl of isolated thylakoids were resuspended in 100 µl of extraction buffer made of 20 mM tricine and 1 mM of phenylmethylsulfonyl fluoride (PMSF). Then the solution was incubated at room temperature for 10 minutes. After the incubation, two freeze/thaw cycles (with vortex) were performed before collecting protein samples by centrifugation (at 3000 xg for 5 minutes) (Lennon et al., 2003). The pellet was collected and resuspended in 100 µl of extraction buffer.
4.3.6. Measuring protein content of the membrane solution using Bradford dye
200 µl of Bradford dye (Bio-rad), 790 µl sterile water and 10 µl of protein extract were mixed in a cuvette. A cuvette with 200 µl of Bradford dye and 800 µl sterile water was used as blank. Solutions were incubated for 10 minutes at room temperature before measuring the protein concentration at
209 595 nm using the Bradford programme in an Eppendorf Biophotometer (Eppendorf, Germany).
4.3.7. Sample preparation from the protein extraction
The same volume of loading buffer (x1) made from Tris-HCl (50 mM, pH 6.8), SDS (2% w/v), glycerol (10% v/v), DTT (100 mM) and Bromophenol blue (0.1% w/v) was added to the protein sample and mixed in an eppendorf tube. Then the sample was denatured by boiling for 5 minutes in a heat block and centrifuged at speed of 12, 000 xg for 10 minutes at 4 oC in a microcentrifuge.
4.3.8. Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE)
Pre-cast SDS gels (4-20% Precise Tris-Glycine gels, Thermo Scientific) were clamped to the electrophoresis tank. Both upper and lower chambers were filled with running buffer (x1) made from Tris-HCl (25 mM), Glycine (200 mM) and SDS (0.1%, w/v). 5 μl of protein samples (10 μg of proteins per slot) were loaded on a pre-cast SDS gel and separated at 150V for 1 hour. After that,
SDS gel were removed from the plastic cassette, placed in a container with a lid and rinsed with distilled water before staining or blotting.
4.3.9. Staining and de-staining of SDS-PAGE
Rinced SDS gels were covered with enough Coomassie Stain (0.1% Coomassie R250, 10% glacial acetic acid and 40% methanol) to cover the gel. Gels were incubated for at least 1 hour on a rocking table before destaining.
Coomassie Stain was poured off from the container and the gel was rinsed with distilled water twice
210 before adding fresh destain solution (20% methanol, 10% acetic acid). Gels were incubated in destain solution on a rocking table overnight.
4.3.10. Measuring chlorophyll content of the thylakoid sample
100 μl of isolated thylakoid suspension was pipetted out and mixed with 900 μl of acetone of 80%
(v/v) and centrifuged (3,000 xg) for 5 minutes. The supernatant was transferred to a glass cuvette and measured the absorbance at 646 or 663 and 750 nm. The following equation was used to calculate the chlorophyll content (µg/ml) in the thylakoid suspension.
Total chlorophyll (Chl a + Chl b) = 17.76 x (A663 - A750) + 7.34 x (A646 - A750) (Porra, et al., 1989)
Where A663 - A750 and A646 - A750 is the difference in absorbance measured at 646 or 663 and 750 nm.
4.3.11. Blue-native polyacrylamide gel electrophoresis (BN-PAGE)
100 µl of isolated thylakoid suspension was centrifuged at 5000 xg for 5 minutes and re-suspended in 75 µl of sample buffer (ACA buffer) made from aminocaproic acid (750mM), Bis-Tris (50 mM, pH 7.0) and EDTA (0.5 mM). Sample preparation was performed at low temperature (4 oC).
Membrane proteins were solubilized by adding 25 µl of n-dodecylmaltoside (0.5-4% w/v) and incubated for 10 minutes at room temperature. After that, the solution was centrifuged at maximum speed (13000 xg) for 30 minutes and the supernatant was carefully removed. 5 ml of Coomassie blue loading buffer made from Coomassie Blue G250 (5% w/v) and aminocaproic acid (750 mM) was added to the supernatant before loading on a gel.
211 20 µl of samples were loaded on a pre-cast blue-native gel (4-16% NativePAGE Novex Bis-Tris gel, Invitrogen by Life Technologies) and separated at 150 V constant voltage for 60 minutes, then voltage was increased to 250 V for 45 minutes at 4 oC (Kügler et al., 1997; Reisinger and
Eichacker, 2006; Breyton et al., 2006).
4.3.12. Western- blot analysis and antibody detection
Western transfer buffer (x1) made from Tris-HCl (25 mM), glycine (192 mM) and methanol (20%) was poured in to a tray. SDS gel, nitrocellulose transfer membrane (Whatman, PROTRAN), two sponges and two stacks of filter papers were wetted using the transfer buffer. The white side of the transfer cassette (anode) was placed on the tray with transfer buffer. Then one of the wet sponges was places on the transfer cassette followed by filter papers. Nitrocellulose membrane was placed on top of the paper towels and then the gel. After that, the other stack of wetted filter papers was placed on the gel followed by the wet sponge. The gel-membrane sandwich was carefully placed in the holder and the chamber was filled with transfer buffer (2/3 full) with a stirrer in the base.
Proteins were transferred from SDS gel to nitrocellulose membrane in cold room (4 oC) at 100 V for
50 minutes to 1 hour.
After transfer the nitrocellulose membrane was stained using Ponceau S stain for approximately 10 minutes to check whether proteins were transferred to the membrane and destained using distilled water. The membrane was blocked using 20 ml of blocking solution made of fresh washing buffer known as Tris buffered Saline (TBS) (150 mM NaCl and 50 mM Tris-HCl pH 8) and Bovine serum albumin (3% w/v) (BSA) and incubated for 1 hour. Polyclonal antibodies were used against PTOX
(provided by Dr M. Kuntz, Universite` Joseph Fourier, Grenoble, France). Commercially prepared antibodies were used for cytochrome f and PsbA (Agrisera, Sweden) and incubated overnight at 4
212 oC. After that, the membrane was washed using TBS and Tween (0.1% v/v) several times. Then a secondary antibody, conjugated with horseradish peroxidase (HRP) was added to the membrane and incubated for 2 hours. Next, the blot was washed several times with TBS and immunoblot detection was performed using ECL western blot developing kit (Amersham, GE Healthcare).
4.3.13. Protein Identification using Mass spectroscopy
Proteins separated on BN-PAGE were identified using mass spectroscopy. Protein samples were sent to the Protein Mass spectrometry core facility centre in the University of Manchester for the identification. Identification have been made using Mascot (version 2.2.06). The statistical validation of the results were performed using Scaffold (version 3.0.04).
4.3.14. Reverse transcription polymerase chain reaction (rt-PCR) using two-step protocol
mRNA from leaves was extracted using Rneasy plant Mini kit (QIAGEN N.V.). Reverse transcription was performed using Bioscript Reverse Transcription kit (Bioline Reagents Ltd, UK).
Primers were designed using Primer3 version 0.4.0 (Whitehead Institute for Biomedical Research)
(Rozen and Skaletsky, 2000). To check the specificity of primers, they were BLASTed against genomic databases available at Phytozome v9.1 (Joint Genome Institute, University of California
Regents, Center for Integrative Genomics). Primers were purchased from Eurogentec (Liege,
Belgium). PCR was performed using a BIO-RAD T100 Thermal cycler. Expression levels were measured relative to the housekeeping gene actin from Thellungiella salsuginea. Table 4.1 contains the primers used rt-PCR analysis.
213 Table 4.1. PTOX and actin primers used in rt-PCR
PTOX Forward 5'-GCCTTTGATTGCTCTTCGAC-3'
Reverse 5'-TCGCTTGAACTCGATGAATG-3'
Actin Forward 5'-ATCGTCCTCAGTGGTGGTTC-3'
Reverse 5'-GGTGCAACCACCTTGATCTT-3'
214 4.4. Results
4.4.1. Plants exposed to salt, drought, cold, different growth irradiances and cold combined with high light
T. salsuginea plants grown for 7 weeks under conditions of 140 µmol m-2 s-1 light intensity and with the day time temperature was 20 oC and night time temperature was 15 oC (controlled conditions) have rosettes of leaves with long petioles (2 cm) and serrated leaf margins (Figure 4.1.a). Plants consist of leaves which are 1-1.5 cm long and 0.5 cm wide. T. salsuginea plants have slow initial growth rate compared to A. thaliana (Taji et al., 2004; Gong et al., 2005; Kant et al., 2006).
Therefore, plants were left for 7 weeks to grow and mature before being used for the stress treatments. In T. salsuginea plants, new leaves initiate from the centre of the rosette. Figure 4.1.b shows 7-week old plants treated with 250 mM of NaCl for 14 days. Some leaves showed chlorosis and decreased leaf area. Figure 4.1.c showed 7-week old T. salsuginea plants under water deficit condition. Old leaves (leaves emerged before the treatment) showed wilting and yellowing, whereas, new leaves did not show any clear indication of stress. However old and new leaves seems to have a thicker waxy cuticle compared to the control plants in Figure 4.1.a. 7-week old plants shown in Figure 4.1.d were grown in low light (100 μmol m-2 s-1) and then transferred to moderate light of 400 μmol m-2 s-1 for 14 days. Plants showed a decrease in leaf area and with thick shiny waxy cuticles. Apart from this, common visible signs of stress, such as leaf drop chlorosis and necrosis were not observed. 7-week old plants exposed to cold (4 oC) and high light (1000 μmol m-2 s-1) for 14 days showed slower growth compared to the control plants (Figure 4.1.e). However, 7- week old plants exposed to cold (4 oC) only for 14 days did not show any visible signs of stress
(Figure 4.1.f).
215 a b
c d
e f
Figure 4.1. Images showing the physical changes of leaves of 7-week old T. salsuginea when exposed to stresses. (a) control plants (b) plants treated with 250 mM of NaCl for 14 days (c) plants under drought for 14 days (d) plants exposed to moderate light (400 μmol m-2 s-1) for 14 days (e) plants treated with cold and high light (4 oC and 1000 μmol m-2 s-1) for 14 days (f) plants treated with cold (4 oC) for 14 days.
216 4.4.2. Immunoblot analysis of PTOX, cytochrome b6f (cytochrome f), PSII (PsbA D1)
Immunoblot analyses were performed using antibodies raised against PTOX, cytochrome f (Cyt f) and PsbA proteins in T. salsuginea (proteins loaded on the basis of equal protein content).
Commercially prepared monoclonal antibodies which are specific cytochrome f and PsbA were used to detect these two proteins on the blot. Polyclonal antibody were used to detect PTOX protein in the membrane. The results of this clearly indicated that exposure of T. salsuginea to drought, salt and different growth irradiances caused an increase in the relative band intensity of PTOX protein, compared to the control plants (Figure 4.2). Compared to control there is no substantial change in the relative band intensity in cytochrome f (Cyt f) and PsbA when exposed to salt, drought or different light intensities.
217 a Control Salt Control Control Drought Dif.L Molecular weight PTOX 40 KDa
Cyt f 32 KDa
PsbA 30 KDa
b
Figure 4.2. (a) Immunoblots and (b) relative band intensity of PTOX, cytochrome f (Cyt f) and
PsbA from control, salt-treated, droughted and plants exposed to different growth irradiance (Dif.
L). Relative band intensity was measured using ImageJ software version 1.46. Each bar in the graph represents the average band intensity for a duplicate sample. Only one band could be observed in each case and that the molecular weight of these bands corresponds to the expected molecular weight of the proteins of interest.
218 4.4.3. Detection and quantification of PTOX gene transcript
Reverse transcription PCR was performed to examine the expression of the PTOX gene transcript in control and salt treated T. salsuginea plants. Figure 4.3 shows the relative expression level of PTOX mRNA in control and salt-treated T. salsuginea plants. It showed the expression of the PTOX gene at different times after salt-treatment. The expression was rapidly induced in response to salt, within the first 3 days. Levels of expression were then maintained throughout the experimental period.
Genomic DNA, contaminating RNA preparations can serve as a template in PCR to produce a false positive. To avoid the contamination of genomic DNA, we followed several methods. Primers are designed using the cDNA sequence of PTOX to avoid the interaction of introns which make large amplified products (larger than the expected cDNA products). In this study we used a filter-based
RNA isolation method and treated with DNase directly on the filter. Then we used 'minus RT'
(mock reverse transcription reaction that did not contain reverse transcription) controls to check the contamination of genomic DNA in samples.
219
a
Hyperladder 1V 1000 900 800 700 600 500 400 300
200
100
c 3d 5d 7d 9d 11d 13d c 3d 5d 7d 9d 11d 13d actin ptox
b
e 3 n e g
X O T P
f
o 2
n o i s s e r p x
e 1
e v i t a l e R 0 control 3 day 5 day 7 day 9 day 11 day 13 day
Figure 4.3. (a) PTOX gene expression along with actin (b) Relative expression level of PTOX mRNA in control and salt-treated plants. It showed the expression of PTOX gene, at different times after salt-treatment. Size of the PTOX band is 125 bps and actin is 107 bps. Data point represent the means of three independent rt-PCR experiments. Abundance of the target RNA was estimated relative to the reference gene actin and data were normalised to the control plants.
220 4.4.4. Analysis of the activity of plastid terminal oxidase (PTOX) when plants exposed to different types of stress conditions.
T. salsuginea plants were illuminated at a range of irradiances and then the efficiency of PSII
(ΦPSII) was measured in the presences of atmospheric O2 and saturated CO2 concentrations (>1200
μL L-1). Plants exposed to salt (250 mM), drought or cold combined with high light (4 oC, 1000
-2 -1 μmol m s ), showed an increased ΦPSII under 20% O2 concentration, relative to the control
(Figure 4.4). However, at 2% O2, ΦPSII decreased when plants were stressed. Plants were grown in different growth irradiances (first grown in low light of 100 μmol m-2 s-1 and then transferred to moderate light of 400 μmol m-2 s-1) and plants grown in greenhouse under semi-natural conditions also showed an increase in ΦPSII at different light intensities. However, this effect on the efficiency of PSII was not observed when plants were exposed to 4 oC.
Abiotic stress including salt, drought, cold combined with high light resulted in increase in electron transport rate of PSII (PSII ETR) in T. salsuginea relative to the control plants. Figure 4.5. showed that this increase was entirely suppressed when the O2 concentration was lowered to 2%. Plants transferred between two different light intensities (100 μmol m-2 s-1 and 400 μmol m-2 s-1) and plants
o grown in greenhouse also showed a sensitivity of PSII ETR to O2. However, plants exposed to 4 C at growth irradiance showed no significant change in ETR between O2 concentrations. PSII ETR in plants increased with light and became saturated at high light intensities. However, PSII ETR of plants grown in the greenhouse and plants treated with cold and highlight (4 oC, 1000 μmol m-2 s-1) was not saturated with light (Figure 4.5 c, f).
221 Salt Cold a b
d c Drought Greenhouse
e Different growth light f Cold and highlight
Figure 4.4. Change in the efficiency of PSII (ΦPSII) measured in control plants (closed symbols) and stressed plants (open symbols) at the different light intensities and CO2 concentration of >1200
-1 μL L . The efficiency of PSII measure at two different O2 concentrations, 2% O2 (black squares)
o and 20% O2 (red circles). (a) Plants treated with 250 mM salt (b) plants treated with 4 C cold (c) plants grew in the greenhouse (d) plants exposed to drought (e) plants exposed to different light intensities (different growth irradiance of 100 and 400 μmol m-2 s-1) and (f) plants exposed to high light (1000 μmol m-2 s-1) and cold (4 oC). Error bars represent the standard error of at least 5 replicates.
222 a Salt b Cold
c d Greenhouse Drought
e f Different growth light Cold and high light
Figure 4.5. Change in the electron transport rate of PSII (ETR of PSII) measured in control plants
(closed symbols) and stressed plants (open symbols) at the different light intensities and CO2
-1 concentration of >1200 μLL . ETR of PSII measure at two different O2 concentrations, 2% O2
(black squares) and 20% O2 (red circles). (a) Plants treated with 250 mM salt (b) plants treated with
4 oC cold (c) plants grew in the greenhouse (d) plants exposed to drought (e) plants exposed to different light intensities (different growth irradiance of 100 and 400 μmol m-2 s-1) and (f) plants exposed to high light (1000 μmol m-2 s-1) and cold (4 oC). Error bars represent the standard error of at least 5 replicates.
223 4.4.5. Isolation of PTOX protein using Blue-native PAGE and Western-blotting
A possible association of PTOX protein with other complexes was investigated by isolating PTOX protein using blue-native PAGE and immunoblotting using anti-PTOX antibodies. Protein samples were prepared from T. salsuginea leaves treated with salt, drought and different growth irradiances for 14 days. Each sample was run along with a control protein sample. Figure 4.4 (a) shows a blue native gel with separated protein complexes of the thylakoid membranes. The gel was compared with the figures with isolated thylakoids from previous studies (Hashimoto et al., 2003; Ivanov et al., 2012). Figure 4.6 (b) shows an immunoblot with a distinct band in the samples exposed to salt, drought and different growth irradiances compared to the control samples. The band is located in the expected region of PSII and LHCII (light harvesting complex II) on the gel. However, mass spectrometry analysis did not identify any peptides predicted from PTOX protein.
Proteins were identified by mass spectrometry analysis (Table 4.2). The table contains the proteins identified, molecular weight and the number of unique peptides that have been matched to the identified protein in that sample (the greater the number of matches the more certain the identification). According to the results, Rubisco and ATP synthase are the most abundant proteins identified in the sample. In addition, subunits of PSI and PSII were identified as prominent proteins in the sample.
224 a b
salt DT c c c Dif.L c salt c DT c Dif.L
-1048KDa
PSII/LHCII -720
-480 PSI/LHCI Rbs PSII,ATPase -242
Cyt b6f dimer -146 LHCII -66 LHCII trimer
Figure 4.6. (a) Blue-native PAGE showing the separated complexes of isolated thylakoids of T. salsuginea (b) Immunoblot analysis showing the presence of prominent signal in control (c), plants exposed to salt, drought (DT) and different growth irradiances (Dif.L). The marked section from the gel was cut and the proteins were identified using the mass spectrometry.
225 Table 4.2. Proteins identified by the mass spectrometry. BN-PAGE separated protein complexes of the thylakoid membranes isolated from T. salsuginea plants exposed to salt, drought and different irradiances. Immunoblot analysis showed a distinct band on the membrane which is specifically located on the region of the PSII and LHCII (light harvesting complex II) on the gel.
Identified Proteins Molecular Weight (KDa) No of unique peptides identified Ribulose bisphosphate carboxylase large chain 53 33 ATP synthase subunit beta 54 24 ATP synthase subunit alpha 55 13 Photosystem I reaction center subunit II-1 23 10 Photosystem II CP47 chlorophyll apoprotein 56 10 Ferredoxin-NADP reductase, leaf isozyme 1 40 9 PsbP-like protein 2 27 8 ATP synthase gamma chain 1 41 6 Photosystem II CP43 chlorophyll apoprotein 52 6 NAD(P)H-quinone oxidoreductase subunit H 46 6 Photosystem I P700 chlorophyll a apoprotein A2 82 5 Photosystem I iron-sulfur center 9 5 Photosystem I reaction center subunit III 24 5 Photosystem I P700 chlorophyll a apoprotein A1 80 4 ATP synthase subunit b 21 4 Photosystem II D2 protein 40 4 Ferredoxin-NADP reductase, leaf isozyme 40 4 Photosystem Q (B) protein 38 3 NAD(P)H-quinone oxidoreductase subunit J 19 3 ATP synthase epsilon chain, 14 3 Malate dehydrogenase, glyoxysomal 37 3 Chlorophyll a-b binding protein 2 28 3 Apocytochrome f 36 3 Photosystem I reaction center subunit V 17 3 Ribulose bisphosphate carboxylase/oxygenase activase 52 2 NAD(P)H-quinone oxidoreductase subunit I 20 2 Geranylgeranyl diphosphate reductase 52 2 Ferredoxin-NADP reductase, leaf isozyme 2 41 2 Photosystem II 22 kDa protein 29 2 Protochlorophyllide reductase B 43 2 Cytochrome b559 subunit alpha 9 2 NAD(P)H-quinone oxidoreductase subunit 2 A 57 2 NAD(P)H-quinone oxidoreductase subunit K, 25 2 Chlorophyll a-b binding protein 8 29 2
226 4.4.6. Genomic information about plastid terminal oxidase
Figure 4.7.a shows the T. salsuginea genomic sequence of PTOX gene (2723 nucleotides) including exons, introns and untranslated regions (UTRs). Figure 4.7.b shows the coding sequence of PTOX with annealing sites for forward and reverse primers used in the rt-PCR analysis. PTOX genome assemblies have only been assembled to the scaffold level and the PTOX gene is located on scaffold_1 (data retrieved from Phytozome v9.1). The schematic representation of the genomic structure of PTOX contains 9 exons which are indicated in yellow coloured boxes. Introns are indicated by lines and transcription start site (ATG) and stop codon (TAA) are indicated.
Untranslated regions (UTRs) 5´-UTR and 3´-UTR are indicated in purple coloured boxes (Figure
4.8.a).
Sequence alignment of PTOX protein from Arabidopsis thaliana (AT) (351 residues) and
Thellungiella salsuginea (TS) (346 residues) showing significant alignment of 91%. PTOX gene and protein sequence from T. salsuginea (Figure 4.8.b). Blast analysis (NCBI, www.ncbi.nlm.nih.go v) of PTOX gene and protein sequence from T. salsuginea showed a sequence similarity (E-values < than 1e-06) to protein IMMUTANS (alternative oxidase family) in A. thaliana (significant alignment, 91% amino acids and 92% nucleotides). The predicted transit peptide of A. thaliana (56 residues) and T. salsuginea (53 residues) are shown in the protein sequence (data retrieved from
ChloroP1.1 Prediction Server, http://www. c bs.dtu.dk/services/ChloroP/ , Uniport protein database,
http:// www . Uni prot. Or g / uniprot and iPSORT Prediction Server, http://ipsort.hgc.jp/predict.cgi). The predicted transmembrane helix domains in both species are indicated by bold letters (21 residues).
Like alternative oxidase (AOX) in mitochondria, PTOX belongs to the family of diiron carboxylate
227 proteins, a group of non-haem iron proteins that contain a coupled binuclear iron centre (Berthold and Stenmark, 2003). Similar to IMMUTANS in A. thaliana, T. salsuginea possess 6 di-iron binding motifs, four glutamates, E136, E175, E227, E296 and two histidine residues, H178 and H299 (A. thaliana IMMUTANS sequence numbering) all of which are completely conserved (McDonald et al., 2011). PTOX protein in T. salsuginea contains AOX (alternative oxidase) and belongs to the
Ferritin-like superfamily, a protein family with four helical bundle domain (Andrews, 2010) (data retrieved from Uniport protein database, http:// www . Uniprot. Org/ uniprot and NCBI Conserved
Domains database, http://www.ncbi . nlm.nih.gov /St r u ctur e /cdd/wrpsb.cg i).
228 a CGATGGGCATGAGAAGGTAGTGGGAGAACCCAGAATATCTGCCCGCCTTGGGAACCGCCTTTCTTCGTATAAGACCCGCA AAATTCCAAAATTTCTCCGTTTCCTACGAAAAATCTCCAACCTTTACTTTTCTTCCCTGTGATCGAATCTTGGGTTCCCT GACGGAGATGGCAGCGACGGTGGCGATTTCAGGCATCTCCCCACGGCCTTTGATTGCTCTTCGACGCTCTAGAGCCGCCG TTTCGTACAGTACTTCTCACCGATTGCTTCTTCATCGTCCTCTCTCTTCTCCTCGCCTGCTCTTCAGGTAGCTACTCGAT TAGAGCACTGAATGATGGAAAACGACAGCTAAATATTTGATTGTTGTCGACTTGTCGTCAAACTTAATGTTGAAAAGGGT TCGATGGGTAGTTTTTTTACTTAGTGCTATCATTTGAACATTTCAGGAACATTCATCGAGTTCAAGCGACGATTTTACAA GACGATGAAGAGAAAGTTGTGGTGGAGGAATCGTTTAAGGCCGAGACTTTTCCTGGTAAAGTACCACTTGAGGAGCCAAA CATGAGTTCTTCAACTAGTGCTCTGGAGGCTTGGATCATCAAGCTTGAGCAAGGAGTGAATGTCTTCCTTACAGTAAGGT TTTATGACCCTTTCTAGGATCTTCAAACAGCTGTGTGTTCAAATTATCCTGTGCATCTAACTAATTGTTGTCTTGTTTTG TTTTTTATCAGGACTCGGTGATTAAGATACTTGACACATTGTACCGTGATCGAACCTATCCTAGGTTCTTTGTTCTTGAA ACAATTGCTAGAGTGCCTTATTTTGGTAAGCGCATCCGTTTGTAGTTTGAGTTCCTCTGATACTCTTGCAAGAAACGTTT AGGGTAGTGTTAAGGATTATTCATTCGGATTTGTGTTCGTCATAAGACAGTAAGAATCGTGAAAGGCACTTGAATTGTGT TAAGAAGAACAATGACAGAATAGTGTTCGGAAGCATATATATACCTTTGTCCATTTCCTTATTTCATTAGAGGCATAATC CATCACCAACCTCAAAGGCCATCTAATCGTACTGTGTAAAATATAGGATTTGTGGAAAATCACATCTCATTTTTTTTCTT GTTGGCTAGAACATAAACATAGTTTATGTATGCTTCTTGCAGCCTTTATGTCAGTGCTACATATGTATGAGACCTTTGGT TGGTGGAGGAGAGCAGATTATTTGAAAGTACATTTTGCTGAGAGCTGGAATGAGATGCACCACTTGCTCATAATGGAAGT AAGAACCACGACTTTCCCTTCTCTACGAGTTTCATACTCTTAACCTAGTAGTTGTGAAAGAGCCAAACGAATGTTTTTGA TGACAGGAATTGGGTGGAAATTCTTGGTGGTTTGATCGTTTTCTAGCCCAGCACATAGCAACCTTTTATTACTTCATGAC GGTGTTCTTGTATATCATAAGCCCGAGGATGGCATGTAGGTTTCATTGACTTCTAGATATTAGCAGAATAAAATCATGAT ATAGAGAAAGGACGACTTCTTGTCTTCATGACCTCATTAACTGTTTGTTACCGTGCAGATCACTTTTCGGAATGTGTTGA GAGTCATGCATATGAGACATACGATAAATTTCTCAAGGCCAGTGGAGGTTGGTTCAACATTTTCAATACTGATTTAGTTT ATCTTTCTCCAACATTTCCTGTCCCAGTTGCATTAGTAATGTAGTTGTTTATGGGTTTAGTGAGCTGAATACCTCACAAA TTCTTCAATGATTTTTACTATCTGGAATGTTTTGCTTCGCTTTTTTTTTTCCCCTGAACAGAGGAGTTGAAGAATTCGCC TGCACCTGATATCGCAGTGAAATACTATACTGGAAGCGACTTGTACTTATTTGGTTAGTTTGTCCTTCCAGTTTTTATCA ATGTTCTCTAGTTCTCAAATTTTCTACCTTTGACTAGTAGGAGTCTCTTTCCGTTTTGTCTGCAGATGAGTTTCAAACAG CTAGAGCTCCCAATACCCGAAGGCCAACAATAGGTACCAAATTAACTTTGTATTATTTCAATTTTTGAATCCATTGAACC ACAAATTACTAAAAAGGTTTATGATTTATCCATGGTTGTGAGTCTACATATTAGGAGGGACTAAGCTTTGGTTTTTCATA CTTATGGATAAAACAGTAATATCAGTTTGTGATTTACTTTGGTGTTGATTTCAGGATCTCTAACGTTGTTGTTGTATTGT GATTATGTGCAGAAAATTTATACGATGTGTTTGTGAACATAAGAGATGACGAAGCAGAACATTGCAAGACAATGAGGGCT TGTCAGACTCTAGGAAGCCTCCGTTCTCCACACTCCATTCTAGAAGACGACGATTGTAATGAAGAATCAGGCTGTGTTGT TCCTGAGGCTCATTGTGAAGGTATTGTAGACTGTATCAAGAAATCCATTACGAATTAAATTAGAACGGAAAAAGGGATTA ATTATATCAACTTATCTTGAAGAATAGATATATCCAATATACTTAGGAATAAAGGAAAGGTGCGAGATTCTCATAGTTAT GTATGTGTGGGGGAGAGAATCAAATACACTTGAGATGTAAAATTATTTTATTCAACCTACTTGATGTTCATCATTGTAGT CGTTTGAGACCATTTTTGTATGCATGCTAGCCATTGTTATTGTATTGCCGGTTTCAATTCATTGGAGTAATATACCAATT TAT b ATGGCAGCGACGGTGGCGATTTCAGGCATCTCCCCACGGCCTTTGATTGCTCTTCGACGCTCTAGAGCCGCCGTTTCGTA CAGTACTTCTCACCGATTGCTTCTTCATCGTCCTCTCTCTTCTCCTCGCCTGCTCTTCAGGAACATTCATCGAGTTCAAG CGACGATTTTACAAGACGATGAAGAGAAAGTTGTGGTGGAGGAATCGTTTAAGGCCGAGACTTTTCCTGGTAAAGTACCA CTTGAGGAGCCAAACATGAGTTCTTCAACTAGTGCTCTGGAGGCTTGGATCATCAAGCTTGAGCAAGGAGTGAATGTCTT CCTTACAGACTCGGTGATTAAGATACTTGACACATTGTACCGTGATCGAACCTATCCTAGGTTCTTTGTTCTTGAAACAA TTGCTAGAGTGCCTTATTTTGCCTTTATGTCAGTGCTACATATGTATGAGACCTTTGGTTGGTGGAGGAGAGCAGATTAT TTGAAAGTACATTTTGCTGAGAGCTGGAATGAGATGCACCACTTGCTCATAATGGAAGAATTGGGTGGAAATTCTTGGTG GTTTGATCGTTTTCTAGCCCAGCACATAGCAACCTTTTATTACTTCATGACGGTGTTCTTGTATATCATAAGCCCGAGGA TGGCATATCACTTTTCGGAATGTGTTGAGAGTCATGCATATGAGACATACGATAAATTTCTCAAGGCCAGTGGAGAGGAG TTGAAGAATTCGCCTGCACCTGATATCGCAGTGAAATACTATACTGGAAGCGACTTGTACTTATTTGATGAGTTTCAAAC AGCTAGAGCTCCCAATACCCGAAGGCCAACAATAGAAAATTTATACGATGTGTTTGTGAACATAAGAGATGACGAAGCAG AACATTGCAAGACAATGAGGGCTTGTCAGACTCTAGGAAGCCTCCGTTCTCCACACTCCATTCTAGAAGACGACGATTGT AATGAAGAATCAGGCTGTGTTGTTCCTGAGGCTCATTGTGAAGGTATTGTAGACTGTATCAAGAAATCCATTACGAATTA A
Figure 4.7. (a) PTOX genomic sequence of T. salsuginea including exons (highlighted in yellow), introns and untranslated regions (UTRs) (highlighted in purple) (b) coding sequence of PTOX which indicates the annealing sites of primers (forward primer highlighted in green and reverse primer highlighted in blue) used in rt-PCR analysis.
229 a ATG TAA
b
AT MA---AISGISSGTLTISRPLVTLRRSRAAVSYSSSHRLLHHLPLSSRRLLLRNNHRVQAT TS MAATVAISGISP------RPLIALRRSRAAVSYSTSHRLLLHRPLSSPRLLFRNIHRVQAT
AT ILQDDEEKVVVEESFKAETSTGTEPLEEPNMSSSSTSAFETWIIKLEQGVNVFLTDSVIKI TS ILQDDEEKVVVEESFKAETFPGKVPLEEPNMSSS-TSALEAWIIKLEQGVNVFLTDSVIKI
AT LDTLYRDRTYARFFVLETIARVPYFAFMSVLHMYETFGWWRRADYLKVHFAESWNEMHHLL TS LDTLYRDRTYPRFFVLETIARVPYFAFMSVLHMYETFGWWRRADYLKVHFAESWNEMHHLL
AT IMEELGGNSWWFDRFLAQHIATFYYFMTVFLYILSPRMAYHFSECVESHAYETYDKFLKAS TS IMEELGGNSWWFDRFLAQHIATFYYFMTVFLYIISPRMAYHFSECVESHAYETYDKFLKAS
AT GEELKNMPAPDIAVKYYTGGDLYLFDEFQTSRTPNTRRPVIENLYDVFVNIRDDEAEHCKT TS GEELKNSPAPDIAVKYYTGSDLYLFDEFQTARAPNTRRPTIENLYDVFVNIRDDEAEHCKT
AT MRACQTLGSLRSPHSILEDDDTEEESGCVVPEEAHCEGIVDCLKKSITS TS MRACQTLGSLRSPHSILEDDDCNEESGCVVP-EAHCEGIVDCIKKSITN
Figure 4.8. (a) The schematic representation of the genomic structure of PTOX contains exons
(yellow coloured boxes), introns (indicated by lines) and transcription start site (ATG) and stop codon (TAA) are indicated. Untranslated regions (UTRs) 5´-UTR and 3´-UTR are indicated in purple coloured boxes. (b) A sequence alignment of PTOX proteins from Arabidopsis thaliana (AT)
(351 residues) and Thellungiella salsuginea (TS) (346 residues) showing significant alignment of
91%. Predicted transit peptide of A. thaliana (AT) (56 residues) is highlighted in green and transit peptide for T. salsuginea (53 residues) is highlighted in yellow. Differences in amino acids in two transit peptides were highlighted in in pink. The amino acids required for iron binding in both sequences are highlighted in yellow. Bold letters indicated the sequences of predicted transmembrane helix domains.
230 4.5. Discussion
As an extremophile, T. salsuginea shows a remarkable potential to thrive in adverse environments.
However, T. salsuginea plants show some physical changes, such as decreased leaf area, slow growth, leaf chlorosis and waxy epicuticular layer (Figure 4.1). The densely distributed epicuticular waxes in leaves of T. salsuginea help plants to protect themselves from high radiation and control water loss (Amtmann, 2009). Studies have shown that T. salsuginea contains low concentrations of
Na+ and Cl- in shoots and has high K+ and Na+ ratio when plants exposed to salt (Inan, 2004; Stepien and Johnson, 2009). It has been found that, compared to A. thaliana, T. salsuginea has an improved ion homoeostasis mechanism and regulates ion transporters under salinity (Inan, 2004). T. salsuginea plants restrict ion toxicity in leaves by ion compartmentalization in the vacuoles which permit leaf initiation and expansion during the stress (M’rah et al., 2006). T. salsuginea accumulates more proline, which acts as a compatible solute and a osmoprotectant compared to A. thaliana
(Inan, 2004; Taji et al., 2004; M’rah et al., 2006; Ghars et al., 2008; Amtmann, 2009). However, studies have shown conflicting results of the differences of proline accumulation in these two species (Inan, 2004; Taji et al., 2004; M’rah et al., 2006; Ghars et al., 2008; Amtmann, 2009). T. salsuginea showed high levels of thioredoxin, which is an important component involved in the defence system against oxidative damage when exposed to stress (M’rah et al., 2006). T. salsuginea can withstand water deficit conditions by maintaining water content and reducing water loss through stomata. Plants decrease the shoot growth under drought and by this protect the shoot meristem from desiccation (Amtmann, 2009).
Work from Stepien and Johnson (2009) showed an increase in the relative abundance of PTOX protein when T. salsuginea plants were challenged with 250 mM of salt. Consistent with that, the immunoblot data in this study demonstrates the upregulation of PTOX protein abundance in T.
231 salsuginea when exposed to salt, drought and different growth irradiances (Figure 4.2). Similar effects were observed in several plants when exposed to extreme environmental conditions. Studies have shown a relatively high amount of PTOX protein in the high mountain species Rananculus glacialis under high light conditions and a decline in activity when plants deacclimated (Streb et al.,
2005; Laureau et al., 2013). Another study by Ivanov and co-workers (2012) showed cold acclimated A. thaliana produces PTOX under high light, although no evidence for significant activity has been shown. Brassica fruticulosa, which is tolerant to heat and high light intensities showed increased levels of PTOX compared to Brassica oleracea (Díaz et al., 2007).
The nuclear encoded gene for PTOX (39-40 kDa) was found by Wu and co-workers (1999) in A. thaliana. PTOX is encoded by single gene in higher plants and two genes (PTOX1 and PTOX2) in some eukaryotic algae (Wang et al., 2009). Results from the mRNA transcripts in this paper ruled out that salt stress caused a steady accumulation of T. salsuginea PTOX gene transcripts and the expression peak was observed after only 3 days of salt treatment (Figure 4.3). Control plants showed the lowest relative expression level, compared to salt treated plants. According to the results, it can be seen that salinity increases the relative expression of PTOX gene in T. salsuginea plants. A similar effect was reported in the study by Streb et al. (2005). According to that study, R. glacialis also showed an increase level of mRNA transcripts in leaves compared to other alpine plants. Work from Simkin et al. (2008) showed an upregulation of PTOX gene transcript in Coffee leaves (Coffea arabica and Coffea canephora) when exposed to drought. OsIM1 gene in a salt tolerant rice mutant, M-20 showed differential expression and produce OsIM1 and pseudo- transcript OsIM2 when exposed to salt (Kong et al., 2003). The amino acid sequence of OsIM1 showed 66% identity with PTOX from tomato. An upregulation of OsIM1 was also found upon exposure to salt and abscisic acid. Haematococcus pluvialis, a unicellular green microalga contains two PTOX genes, PTOX1 and PTOX2 (Li et al., 2008; Li et al., 2010). This green alga showed an
232 increased in both gene expression when exposed high light, sodium acetate and ferrous sulfate. A study with Chlamydomonas reinhardtii showed an increase in both PTOX1 and PTOX2 gene transcripts under phosphorous limitation (Moseley et al., 2006). However, a study Houille-Vernes et al. (2011) found that PTOX2 is the major oxidase involved in chlororespiration in Chlamydomonas reinhardtii. The PTOX gene expression pattern of T. salsuginea observed in this study is different than the PTOX activity pattern observed in the study performed by Stepien and Johnson (2009), where the activity was induced more slowly. This shows that the PTOX transcript is rapidly induced and that the protein accumulates more slowly. This could imply two levels of control that operate on different time scales.
PTOX activity was measured by comparing electron transport rates at two O2 concentrations. In the alternative pathway, excess electrons are transferred to O2 to produce water, therefore, decreases in the O2 concentration can reduce the activity of PTOX, which can be seen as a drop in ΦPSII.
According to the results, in T. salsuginea in addition to salinity, drought, different growth irradiances and cold and high light together also trigger the production of the plastid terminal oxidase which is actively involved in increasing the efficiency of PSII (Figure 4.4). Plants grown in unheated greenhouse also showed an activity of PTOX (Figure 4.4.c). This suggests that PTOX expression is, to a greater or lesser extent, a normal response of T. salsuginea and not a response to extreme stress. However, plants grown in 4 oC did not show significant activity of PTOX. Griffith et al. (2007) found out that T. salsuginea plants are resistance to freezing temperatures from -13 to
18.5 oC and can complete their life cycle at 5 oC. Therefore, exposing T. salsuginea plants to 4 oC may not be severe enough to trigger the activity of PTOX and it may only harden the plants to withstand freezing conditions. This is clear, because plants showed an activity of PTOX when cold stress was combined with high light (4 oC and 1000 μmol m-2 s-1). This consistent with a study showing that PTOX is not induced when Ranunculus glacialis was exposed to low temperature but
233 was induced when exposed to high light and low temperature (Laureau et al., 2013). At ambient O2 concentration, PSII ETR of plants grown in a greenhouse and plants treated with cold and high light
(4 oC and 1000 μmol m-2 s-1) was not saturated with light (Figure 4.5.c and Figure 4.5.f). This suggested that, T. salsuginea acclimated to changing environmental conditions. Previous studies have suggested that PTOX could play a role in the acclimation of photosynthesis under changing environmental conditions (Peltier et al., 2002; Kuntz, 2004; Rumeau et al., 2007; Díaz et al., 2007;
Trouillard et al, 2012).
Multiple alignment and blast analysis showed that the PTOX in T. salsuginea showed a sequence similarity to the PTOX protein in A. thaliana. Similar to that, PTOX in T. salsuginea contains an N- terminal chloroplast targeting sequence (putative transit peptide) (Carol et al., 1999; Wu et al.,
1999; McDonald et al., 2011). The iPSORT software (Nakai and Kanehisa, 1992) predicted the first
53 amino acids of the protein sequence contain N-terminal transit sequence which targets the chloroplast. PTOX and AOX belong to the non-haem diiron carboxylate protein family (DOX)
(Berthold et al., 2002, Moore et al., 2008; Shiba et al., 2013). All the proposed structures and the recently published crystal structure from Trypanosoma brucei, AOX is a homodimer with non-haem diiron carboxylate active site within a four helix bundle (Andersson and Nordlund, 1999; Berthold et al., 2000; Berthold et al., 2002, Moore et al., 2008; Shiba et al., 2013). This helix bundle provides six ligands to bind a diiron center which are conserved in all AOX and PTOX proteins. They are four glutamate (E136, E175, E227, E296) and two histidine residues (H178, H299) (A. thaliana
IMMUTANS sequence numbering) (Berthold et al., 2002; Moore et al., 2008; McDonald et al.,
2011). These ligands are essential for the activity of DOX proteins and do not tolerate change (Fu et al., 2005). A study from Shiba et al. (2013) showed that histidine residues which are distant from diiron center form H bonds with other residues to build a network. This is important as it stabilize the active site of AOX. Although AOX is a homodimer (Shiba et al., 2013), a recent study by Yu et
234 al. (2014) showed that PTOX protein in rice exists mainly as a homo-tetrameric complex. This study has shown that tetrameric complex of PTOX contained two Fe per monomer which is similar to the motifs present in the primary structure and with the crystal structure obtained with the mitochondrial AOX as observed in Shiba et al. (2013).
PTOX plays a major role as an electron sink during photosynthesis in Chlamydomonas reinhardtii mutants lacking either PSI or Cyt b6f (Cournac et al., 2000). A study has shown that PTOX plays an important role in lowering the over-reduction of PSII in the gun4 mutant of Chlamydomonas reinhardtii where electron transport from PSII to PSI is strongly decreased (Formighieri et al.,
2012). They also showed that an increase in PSII excitation pressure when treating this mutant with propylgallate, an inhibitor of PTOX. In addition, a study showed that Synechococcus WH8102, a marine cyanobacterium possess an alternative electron flow to O2 via PTOX when PSI activity is limited due to low iron levels. They hypothesized that Synechococcus uses PTOX which only have two iron atoms rather cytochrome b6f and PSI which have 18 iron atoms altogether to survive in low iron conditions (Bailey et al., 2008; Cardol et al., 2008). These studies suggested a direct electron transfer from PSII to oxygen via the PQ pool to produce water under stress conditions.
Treatment with PTOX inhibitor, n-propyl gallate and the cytochrome b6f inhibitor, 2' iodo-6- isopropyl-3-methyl-2',4,4'-trinitrodiphenylether (DNP-INT) suggested the involvement of PTOX in direct electron transfer from PSII to oxygen to control PQ redox state in T. salsuginea under salt stress (Stepien and Johnson, 2009). In addition, it has been suggested that, PTOX may regulate in the NDH-dependent cyclic pathway under abiotic stress (Rumeau et al., 2007). Previous studies on
PTOX by Kuntz (2004) showed an involvement of NDH in directing electrons from the PQ pool to
PTOX and then to molecular oxygen. A study by Streb and co-workers (2005) showed an increase in NDH with increasing PTOX in high mountain species Rananculus glacialis, suggesting an involvement in NDH-dependent cyclic electron transport. Cold acclimated A. thaliana produced
235 high PTOX protein content but low NDH suggesting the absence of an NDH-dependent cyclic electron transport pathway (Ivanov et al., 2012). Increased NDH and PTOX levels in Brassica fruticulosa than Brassica oleracea suggested, the involvement of chlororespiratory process in adaptation to heat and high illumination (Díaz et al., 2007).
According to predicted pathways, PTOX is assumed to interact with the PQ pool independently from other thylakoid complexes. However, similar to alternative oxidase in mitochondria (Navet et al., 2004; Duarte and Videira, 2009), there is a high possibility that PTOX is associated with one of the other electron transport complexes, most probably cytochrome b6f or PSII. Due to the high protein concentration, diffusion of plastoquinol in the thylakoid membranes is restricted (Kirchhoff et al., 2008). Therefore, for efficient electron transport, PTOX would be located near PSII (Stepien and Johnson, 2009). PSII is located mainly in the granal stacks (Albertsson, 2001). However,
Lennon et al. (2003) found that PTOX is localized in the stromal lamellae in spinach. PTOX is known to act an electron sink which inhibits the production of ROS in many plants and alga under environmental stresses. However, several studies showed that over-expressing PTOX protein in other photosynthetic organisms promotes photo-oxidative damage and causes photoinhibition compared to the wild type (Heyno et al., 2009; Ahmad et al., 2012). Therefore, the protective function of PTOX may not be universal to all the photosynthetic organisms in the world. High amounts of PTOX and prominent upregulation of photosynthesis even under severe stress conditions suggested that some plants, including T. salsuginea, R. glacialis and algae including,
Haematococcus pluvialis have an effective stress tolerant strategy different to other photosynthetic organisms. A study by Yu et al. (2014) showed that, PTOX in rice can act as a safety valve when the steady state PQH2 is low, while a certain amount of ROS is formed at high light intensities. In this study, they also suggested that, the structure of PTOX might change with the plant species and the possible association with cytochrome b6f would control the electron flow from plastoquinol to
236 PTOX. Immunoblot analyses were performed to assess any change in relative abundance of these complexes when exposed to salt, drought and high light. The results suggested that there is no substantial change in the relative abundance of cytochrome b6f and PSII complexes when PTOX is expressed under stress. However, further analysis is needed to identify whether the PTOX is in appressed regions in T. salsuginea.
Understanding the activity of PTOX under stress conditions, including possible pathways in the electron transport chain and its location in the thylakoid membrane may open the possibility of engineering PTOX into stress sensitive plants. Figure 4.6 showed the experiment performed to identify the location of PTOX and the other associating complexes. The prominent signal on immunoblot located in the region of PSII and LHCII in the blue native PAGE gel. According to the results, plants treated with salt, drought and different growth irradiances showed more intense signals compared to the control plants. A number of proteins were identified and the most abundant proteins are subunits of Rubisco, which is a common contaminant of thylakoid preparations. In addition, subunits of PSII and PSI were identified. Presence of PSII subunits supports the assumption of the association of PTOX with PSII supercomplex as this would place PTOX in the same location as the reduced PQ pool. However, the mass spectrometry analysis did not show any indication of PTOX protein which we cannot support the assumption precisely. The region where the band was seen and the presence of many different complexes suggests possibly that this is not a region with a high concentration of any one complex, so there is no clear evidence for a specific association. Therefore, more detailed analysis is needed to discover the precise location of PTOX and any associated complexes in T. salsuginea. Two-dimensional gel electrophoresis would be an ideal technique to use to separate thylakoid proteins. This technique can be use where high resolution separation of proteins is needed (O'Farrell, 1975). Apart from that, other fractionation techniques, such as separating grana from stroma using digitonin preparations of sonication
237 followed by aqueous two phase separation can be used (Stefánsson et al., 1997).
In conclusion, we have shown a substantial increase in relative abundance in PTOX protein in T. salsuginea plants exposed to salt, drought and different growth irradiances compared to the control plants. Apart from that, we observed an upregulation of PTOX gene transcripts in T. salsuginea plants under salt stress and induced activity of PTOX when exposed to salt, drought, different growth irradiances, cold combined with high light and plants grown in greenhouse conditions.
These results suggested that PTOX is involved in the photoprotection of T. salsuginea plants under abiotic stress conditions. Direct electron transport from PSII to PTOX then to oxygen via the PQ pool is accounting for up to 30% of total PSII electron flow in T. salsuginea (Stepien and Johnson,
2009). Efficient electron flow from PSII to PTOX would however, probably require co-location of these complexes in the same thylakoid fraction. However, our attempt to identify the precise location of PTOX in the thylakoid membrane was not successful. Therefore, further studies are needed to find the precise location of PTOX protein in T. salsuginea.
238 Chapter 5
General Discussion
239 Abiotic stress including salt, drought and extreme temperatures, causes adverse effects on growth and productivity of plants. According to the Declaration of the World Summit on Food Security, in less than 40 years, the global population is expected to increase above 9 billion (FAO. Declaration of the World Summit on Food Security, 2009; Hirayama and Shinozaki, 2010). As the world population increases, it is essential to develop crops, which give high yield (Caemmerer et al.,
2012). Use of extremophile crops that are adapted to adverse environmental conditions might be a possible solution to the future food crisis (Bressan et al., 2011). There are several examples for the use of halophytes for industrial, ecological, or agricultural purposes (Koyro et al., 2011). Salicornia bigelovii is an oilseed halophyte which has 28% oil and 31% protein and similar to soybean yield and seed quality (Glenn et al., 1999). Another example is Panicum turgidum, a halophytic grass which is use as an animal fodder and studies have shown that animals fed with this grass provided better quality meat with high protein and low fat content (Koyro et al., 2011). Yield potential of crops is often limited by the photosynthetic capacity, especially under stress. Therefore, a better understanding of stress tolerance mechanisms in plants will help us to cultivate marginal lands and increase the food production for the growing population (Wu et al., 2012). The concept called cash crop halophytes was introduced few years ago to identify and select plant species which are tolerant to salt stress (Koyro et al., 2011). In this technique, plants are irrigated with saline water and then screening methods are applied to identify salt resistant plants and use biomarkers to characterise halophytes (Koyro et al., 2011). However, in order to successfully apply this method, it is essential to understand the salt tolerance mechanisms in plants, especially crops. The evaluation of the physiology of photosynthesis in stress tolerant and stress sensitive plants started in this work may be important in future attempts to engineer high yielding crops which can grow in adverse environmental conditions.
The study on the regulation of photosynthesis in barley gives a basic understanding of the stress
240 responses and the regulatory processes in photosynthesis under salinity. Barley showed tolerance to salt concentrations higher than 100 mM. However, it showed clear stress responses when exposed to salt as high as 250 mM. We suggested that, at low salt concentrations plants protect PSII centres from excitation pressure by down-regulating the electron transport chain and maintaining a pH gradient across the thylakoid membranes by cyclic electron transport associated with PSI, to support
NPQ. However, at 250 mM this regulation starts to fail. Despite of having more 'active' PSI centres, which suggested the increased cyclic electron transport at 250 mM, a drop of NPQ might be due to the increased leakiness of the thylakoid membranes. Studies of Sharkey (2005) and Sharkey and
Zhang (2010) showed that moderately high temperatures (35-45 oC) induce cyclic electron flow, but proton leakage through the membranes. The membrane leakiness could be measured by using the electrochromic shift (Witt, 1979). This method which uses three wavelengths (505, 520, 535 nm) to exclude interfering signals from light scattering and zeaxanthin, gives the pH component of the proton motive force (Zhang et al., 2009; Sharkey and Zhang, 2010). Apart from that, chlorophyll a/b ratio decreased at 250 mM suggesting a specific loss of reaction centers in both photosystems or loss of PSI compared to PSII. Therefore, further experiments are needed to analyse the effects of salt on light harvesting complexes and reaction centres in two photosystems. This could be performed by examining the changes in protein contents in these complexes using immunoblot analysis. Alternatively, quantitative proteomic mass spectrometry analysis will provide information on the effects of salt on plants through accurate quantification of proteins (Dost et al., 2012;
Liebler and Zimmerman, 2013; Wasinger et al., 2013). In addition, relaxation kinetic studies will provide information about the qE and photoinhibition components in NPQ in the salt-treated barley leaves. In this study, effects of salinity on barley were shown after 14 days of salt treatment.
Because of that, most of the data provided only long term photosynthetic responses of barley to salt rather than both long term and short term salinity responses. Therefore, it is important to extend this study by performing time course experiments, showing both short term and long term responses of
241 barley to salt stress. Salt will not enter leaves immediately, but may change gradually over time, so it is important to monitor leaf salt concentrations at the same time. Apart from that, a comparative study with a salt tolerant barley cultivar (Ligaba and Katsuhara, 2010) or species, such as Hordeum maritimum (Lombardi et al., 2000) will provide important information on the salt tolerance of barley.
Physiological evaluation of salt stress of two rice varieties from Sri Lanka were performed to understand the physiology of photosynthesis in salt-tolerant and salt-sensitive plants and characterize the salt-tolerant traits in plants which are responsible for the regulation of photosynthesis. In this study, we have addressed the effects of salinity on two developmental stages, the early vegetative and the flowering stages of rice, whereas most studies only focus on a one stage of the life cycle. In both stages, At-354, the salt-tolerant variety, is relatively less affected and showed more prominent traits of salinity tolerance than the salt-sensitive Bg-352. This is reflected in photosynthesis of Bg-352 being inhibited at the lower salt concentrations compared to At-354.
However, at 100 mM, the regulation starts to fail even in the salt tolerant At-354. The study of salt stress on barley and rice pointed out that barley is more salt tolerant than rice. Barley is tolerant to salt concentrations higher than 100 mM whereas, even the salt-tolerant rice variety, At-354 is sensitive to salt concentrations higher than 100 mM. However, increasing salinity decreases CO2 assimilation in both barley and rice causing a decrease in photosynthesis. At low salt concentrations this effect is more stomatal in both crops and the effect become non-stomatal with increasing salt concentration. A better understanding of the salt tolerance of barley may allow identification of traits that could be transferred to rice.
The evaluation of the salt-tolerant and salt-sensitive traits started in this work will provide important information on the future attempts to produce salt-tolerant and high yielding rice
242 varieties. Due to the difficulties occurred during the growth periods, this study focused only on the physiological analysis of salt stress regulation in two rice varieties. However, further analysis are needed to examine the change in ion concentrations in leaves of two rice varieties. A biochemical analysis may also be worth doing to understand the effects of salt stress on the thylakoid composition and the Rubisco activity in two rice varieties. Similar to barley, the salt sensitive Bg-
352 showed a drop of NPQ at 100 mM at the early vegetative stage, which might be due to salt induced membrane leakiness and it could be measured by using the electrochromic shift (Witt,
1979). As we have mentioned earlier, it is important to perform a time course experiment to show both short term and long term responses of rice to salt stress. Most of the crops are highly sensitive to abiotic stress during the flowering stage and caused adverse effects on the seed production
(Cominelli et al., 2013). Therefore, examining the effects of salt on the seed production in two rice varieties is also worth doing because it is the most important parameter from an agronomical point of view. The physiological evaluation of both of these crops showed that, at low salt concentrations, plants use cyclic electron transport to regulate electron transport chain. However, this regulation starts to fail with increasing salt concentration. These results suggested that although pathways like cyclic electron transport around PSI act as preferred photoprotective mechanism in plants, it alone may not be sufficient to improve the stress tolerance in sensitive plants. Therefore, it is essential to produce crop varieties with improved regulatory processes to withstand adverse environmental conditions.
The PTOX protein found in Thellungiella salsuginea (T. salsuginea) regulates electron transport by diverting excess electrons in the PQ pool to oxygen to produce water under salinity (Stepien and
Johnson, 2009). Current study has shown that, in addition to salinity, drought, different growth irradiances, greenhouse conditions and cold combined with high light also trigger the production of
PTOX in T. salsuginea which is actively involved in increasing of the efficiency of PSII. The
243 immunoblot data in this study demonstrates the upregulation of PTOX protein abundance in T. salsuginea when exposed to salt, drought and different growth irradiances. However, we were unable to show the results of immunoblot analysis for the plants challenged with cold and highlight and plants grown in greenhouse due to experimental difficulties to collect enough thylakoids from these.
Failure to induce PTOX and activate the alternative pathway in over-expressed transgenic plants suggests that PTOX may be a subunit of some larger thylakoid protein complex, favourably PSII and cytochrome b6f (Heyno et al., 2009; Ahmad et al., 2012). Therefore, immunoblot analysis were performed to identify any change in these two complexes when exposed to abiotic stress. However, results did not show any significant change in the protein levels in these two complexes. Therefore, to fully elucidate interactions of PTOX protein with other thylakoid protein complexes and the precise location on the thylakoid membrane, further analysis needs to be done. Blue-native PAGE and the immunoblot analysis were performed to identify the associated protein complexes which will provide the specific location of the PTOX protein on the thylakoid membrane. Although proteins including, Rubisco, ATP synthase, several subunits of PSII and PSI complexes were identified using the mass spectrometry, we failed to identify PTOX. Therefore, more detailed analysis is needed to discover the precise location of PTOX and any associated complexes in T. salsuginea. Two-dimensional gel electrophoresis would be an ideal technique to use to separate thylakoid proteins. This technique can be use where high resolution separation of proteins is needed
(O'Farrell, 1975). Apart from that, other fractionation techniques, such as separating grana from stroma using digitonin preparations of sonication followed by aqueous two phase separation can be used (Stefánsson et al., 1997). It is possible that loose associations between proteins maybe important and these are less likely to be maintained in native gels. To address these, different cross linkers could be used to identify specific interactions between PTOX and other peptides (Miernyk
244 and Thelen, 2008; Ido et al., 2014).
Although the results presented in here and the study performed by Stepien and Johnson (2009) have demonstrated that, PTOX plays a protective role in T. salsuginea under abiotic stress, it is important to know whether PTOX alone is sufficient to provide the activity as a safety valve in T. salsuginea and whether that activity can be transferred into another plant species. RNA interference (RNAi) is an RNA silencing method which blocks gene function through inserting short sequences of RNA that match part of the target gene’s sequence, therefore, no proteins are produced (Baulcombe,
2000; Matzke et al., 2001; Agrawal et al., 2003; Eamens et al., 2008; Angaji et al., 2010). This technique silences individual genes producing knockout phenotypes, through transformants which produce the required hairpin RNAs or infecting with recombinant RNA viruses that carry the target gene and thus provides information about the function of important genes (Tenea, 2009). RNAi could used to downregulate the PTOX expression in T. salsuginea and, thereby, the involvement of
PTOX as an alternative electron sink can be analysed. This technique was successfully used to examine gene function in many plants, including tobacco, Arabidopsis thaliana, cotton and rice
(Wesley et al., 2001; Stoutjesdijk et al., 2002). Successful attempts of overexpressing Arabidopsis thaliana PTOX in tobacco (Joët et al., 2002), constitutively in A. thaliana (Rosso et al., 2006) and
Chlamydomonas reinhardtii PTOX in tobacco (Ahmad et al., 2012) have been reported earlier.
However, these studies have not lead to a significant increase in PTOX activity under stress.
Therefore, it would be interesting to perform this with the gene from T. salsuginea, to see if the peptide produced has different activity to those from other species. For example, transgenic A. thaliana constitutively overexpressing T. salsuginea PTOX will provide information about role of
PTOX in the photoprotection.
In conclusion, it is thought that the data in this thesis has provided new insights into how the stress
245 sensitive crop plants, such as barley and rice regulate electron transport under salinity and how the regulation fails when expose to high salt concentrations. In addition, this thesis has shown that the alternative electron transport associated with PTOX act as a safety valve under many abiotic stress conditions. Coupling between PTOX activity with other regulatory processes in photosynthesis will help stress sensitive crops to grow and reproduce under changing environmental conditions.
Introducing PTOX gene into stress sensitive crops will be the future goal of this project. However, detailed study on structure and the function of PTOX, possible pathways, enzyme kinetics and the precise location on the thylakoid membrane is necessary when engineering PTOX gene into stress sensitive crops such as rice.
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