STRUCTURAL PHYSIOLOGICAL AND MOLECULAR CHARACTERISATION OF ...eprints.qut.edu.au/90050/1/Mohammad Reza_Karbaschi_Thesis.pdf · Structural, physiological and molecular characterisation

STRUCTURAL, PHYSIOLOGICAL AND

MOLECULAR CHARACTERISATION OF THE

AUSTRALIAN NATIVE RESURRECTION

GRASS TRIPOGON LOLIIFORMIS

(F.MUELL.) C.E.HUBB. DURING

DEHYDRATION AND REHYDRATION

Mohammad Reza Karbaschi

Submitted in fulfilment of the requirements for the degree of

Doctor of Philosophy

Centre for Tropical Crops and Biocommodities

Science and Engineering Faculty

Queensland University of Technology

November 2015

Structural, physiological and molecular characterisation of the Australian native resurrection grass Tripogon

loliiformis (F.Muell.) C.E.Hubb. during dehydration and rehydration i

Keywords

Arabidopsis thaliana; Agrobacterium-mediated transformation; Anatomy;

Anti-apoptotic proteins; BAG4; Escherichia coli; Bulliform cells; C4 photosynthesis;

Cell wall folding; Cell membrane integrity; Chaperone-mediated autophagy;

Chlorophyll fluorescence; Hsc70/Hsp70; Desiccation tolerance, Dehydration;

Drought; Electrolyte leakage; Freehand sectioning; Homoiochlorophyllous; Leaf

structure; Leaf folding; Reactive oxygen species (ROS); Resurrection plant;

Morphology; Monocotyledon; Nicotiana benthamiana; Photosynthesis; Physiology;

Plant tissue; Programed cell death (PCD); Propidium iodide staining; Protein

microarray chip; Sclerenchymatous tissue; Stress; Structure; Tripogon loliiformis;

Ubiquitin; Vacuole fragmentation; Kranz anatomy; XyMS+;

ii Structural, physiological and molecular characterisation of the Australian native resurrection grass Tripogon

loliiformis (F.Muell.) C.E.Hubb. during dehydration and rehydration

Abstract

Plants, as sessile organisms must continually adapt to environmental changes.

Water deficit is one of the major environmental stresses that affects plants. While

most plants can tolerate moderate dehydration (leaf water potential from -5 to -10

MPa) a small group of vascular plants can tolerate desiccation to an air-dry state (-

100 MPa) and beyond. Since the initial discovery of desiccation-tolerance in plants

in 1912 by Irmscher, scientists and plant botanists world-wide have been fascinated

by these unique plants, particularly how these land-plants can tolerate desiccation

and resurrect. Tripogon loliiformis is a largely uncharacterised Australian

desiccation-tolerant grass that resurrects from the desiccated state within 72 hours.

The work performed in this thesis involved a combination of structural, physiological

and molecular techniques to investigate the unique structural, physiological and

molecular features that enable T. loliiformis, and potentially other resurrection plants,

to tolerate desiccation. The molecular studies were performed using high-throughput

protein microarray technology which provided a platform for comparative analysis of

potential protein-protein interacting partners with the pro-survival/anti-apoptotic

protein, Bcl-2 associated athanogene 4 (BAG4). In addition to analysis of the BAG4

protein from Arabidopsis, a novel orthologue of AtBAG4 was isolated from T.

loliiformis and expressed, thus allowing an investigation of the roles of this protein

between desiccation sensitive (A. thaliana) and tolerant (T. loliiformis) plants.

Key observations included; i) a myriad of structural changes such as leaf

folding, cell wall folding and vacuole fragmentation that mitigate desiccation stress,

ii) potential role of sclerenchymatous tissue within rapid leaf folding and light

protection, iii) retention of approximately 70 % chlorophyll in the desiccated state,

iv) early shutdown of photosynthesis, 50 % at 80 % relative water content (RWC)

and ceasing completely at 70 % RWC, v) the possible contribution of bulliform cells

in leaf folding, water reserve and a key role in photosynthesis shut down by

dehydration, vi) a sharp increase in electrolyte leakage during dehydration, vii)

confirmation of membrane integrity by propidium iodide staining throughout

dehydration, desiccation and rehydration, and viii) the molecular demonstration of a

large number of proteins that possibly interact with BAG4 which mostly are related


loliiformis (F.Muell.) C.E.Hubb. during dehydration and rehydration iii

to carbohydrate pathways involved in autophagy. Taken together, these results

demonstrate that T. loliiformis implements a range of structural and physiological

mechanisms, both early on and throughout the drying, that protect tissues from

mechanical, oxidative and irradiation stress. These results confirm that resurrection

plants actively participate in stress tolerance and provide insights into tolerance

mechanisms utilized by these unique land-plants for potential utilization in

enhancement of stress-tolerance in crop plants.

The protein results from protein microarray chip demonstrated that BAG4 has

anti-apoptotic properties due to its interaction with a large number of proteins

involved in autophagy (particularly carbohydrates). Detoxification and recycling of

damaged and unwanted proteins result in recovery of the cells and prevent apoptosis.

More proteins interacted with the TlBAG4 compared with AtBAG4 which might

suggest that the more binding sites exist on BAG4 from this resurrection plant which

in return might contribute in desiccation-tolerance. Although the exact involvement

of BAG4 protein with other proteins and its true involvement in autophagy remained

to be explored, this project for the first time suggested a potential role for BAG4 in

plant autophagy pathways. Furthermore, the data generated from the protein-protein

interaction of BAG4 using high-density protein microarrays provided a valuable

resource for uncovering the mechanisms/pathways that this protein influences for

future research.

iv Structural, physiological and molecular characterisation of the Australian native resurrection grass Tripogon


Table of Contents

Keywords ................................................................................................................................................. i

Abstract ................................................................................................................................................... ii

Table of Contents ................................................................................................................................... iv

List of Figures ....................................................................................................................................... vii

List of Tables ......................................................................................................................................... ix

List of Abbreviations .............................................................................................................................. x

Statement of Original Authorship ........................................................................................................ xiv

Acknowledgements ............................................................................................................................... xv

CHAPTER 1: INTRODUCTION AND LITERATURE REVIEW ................................................. 1

1.1 Introduction.................................................................................................................................. 1

1.2 Classification of land Plants based on their tolerance toward water deficit ................................. 2

1.3 Desiccation tolerance in plants .................................................................................................... 3 1.3.1 Evolutionary aspects ......................................................................................................... 3 1.3.2 Geographic distribution and ecology ................................................................................ 3 1.3.3 Desiccation-tolerant plants types ...................................................................................... 6

1.4 Impacts of water deficit on plants .............................................................................................. 11 1.4.1 Plant responses to water deficit ...................................................................................... 11 1.4.2 Water deficit response characteristics common among all plants .................................. 12

1.5 Structural aspects ....................................................................................................................... 12 1.5.1 Leaf surface structures .................................................................................................... 12 1.5.2 Reducing leaf surface area .............................................................................................. 13 1.5.3 Xylem tissue (in stem and root) ...................................................................................... 15

1.6 Physiological aspects ................................................................................................................. 16 1.6.1 Reactive oxygen species ................................................................................................. 16 1.6.2 Photosynthesis ................................................................................................................ 17 1.6.3 Respiration ...................................................................................................................... 18

1.7 Molecular responses to water deficit in plants ........................................................................... 19 1.7.1 Molecular responses to water deficit in desiccation-tolerant plants ............................... 19 1.7.2 Regulatory ...................................................................................................................... 20 1.7.3 Antioxidants ................................................................................................................... 21 1.7.4 Carbohydrates level in desiccation tolerant plants in resurrection plants ....................... 22 1.7.5 Proline and proteins ........................................................................................................ 25

1.8 Tripogon loliiformis (F.Muell.) C.E.Hubb. a native Australian resurrection grass .................... 28 1.8.1 Australian resurrection plants ......................................................................................... 28 1.8.2 Tripogon Roem. & Schult a genus of true grasses......................................................... 30 1.8.3 Tripogon loliiformis (F.Muell.) C.E.Hubb an Australian resurrection grass .................. 33

1.9 BAG, a family of pro-survival proteins ..................................................................................... 36 1.9.1 In animals ....................................................................................................................... 37 1.9.2 In plants .......................................................................................................................... 37

1.10 Conclusion, aims and objectives ................................................................................................ 38 1.10.1 Aims ............................................................................................................................... 38 1.10.2 Objectives ....................................................................................................................... 39

CHAPTER 2: GENERAL MATERIALS AND RESEARCH DESIGN........................................ 41

2.1 Research Design ........................................................................................................................ 41 2.1.1 Structural experiments .................................................................................................... 41


loliiformis (F.Muell.) C.E.Hubb. during dehydration and rehydration v

2.1.2 Physiological experiments .............................................................................................. 44 2.1.3 Molecular experiments ................................................................................................... 46

2.2 General Materials ....................................................................................................................... 49 2.2.1 Sources of specialised reagents ....................................................................................... 49 2.2.2 Oligodeoxyribonucleotide synthesis ............................................................................... 49 2.2.3 Bacterial strains .............................................................................................................. 49 2.2.4 List of general solutions .................................................................................................. 49 2.2.5 Plant material .................................................................................................................. 50

CHAPTER 3: TRIPOGON LOLIIFORMIS DISPLAYS STRUCTURAL FEATURES AND

CHANGES THAT PROTECT IT DURING DEHYDRATION ..................................................... 53

3.1 Introduction ................................................................................................................................ 53 3.1.1 Structural features ........................................................................................................... 53 3.1.2 Structural changes ........................................................................................................... 53

3.2 Materials and methods ............................................................................................................... 54 3.2.1 Plant materials ................................................................................................................ 54 3.2.2 Methods .......................................................................................................................... 55

3.3 Results ........................................................................................................................................ 56 3.3.1 General structural observations ...................................................................................... 56 3.3.2 Structural changes ........................................................................................................... 63

3.4 Discussion .................................................................................................................................. 68

CHAPTER 4: PHYSIOLOGICAL RESPONSES OF TRIPOGON LOLIIFORMIS LEAVES

DURING DEHYDRATION AND REHYDRATION ...................................................................... 73

4.1 Introduction ................................................................................................................................ 73

4.2 Materials and methods ............................................................................................................... 74 4.2.1 Plant materials and dehydration and rehydration treatment ............................................ 74 4.2.2 Methods .......................................................................................................................... 74

4.3 Results ........................................................................................................................................ 76 4.3.1 Leaf water status and pigmentation ................................................................................ 76 4.3.2 Chlorophyll a fluorescence ............................................................................................. 77 4.3.3 Estimation of the membrane integrity............................................................................. 80 4.3.4 Photosynthesis rate ......................................................................................................... 83

4.4 Discussion .................................................................................................................................. 85

CHAPTER 5: PROTEIN-PROTEIN INTERACTION PROFILE OF BAG4 SUGGESTS A

POSSIBLE ROLE WITHIN CHAPERONE-MEDIATED AUTOPHAGY .................................. 90

5.1 Introduction ................................................................................................................................ 90 5.1.1 Programmed cell death ................................................................................................... 90 5.1.2 BAG4 .............................................................................................................................. 91 5.1.3 Protein microarray .......................................................................................................... 92

5.2 Materials and methods ............................................................................................................... 93 5.2.1 Polymerase chain reaction (PCR) ................................................................................... 95 5.2.2 Agarose gel electrophoresis ............................................................................................ 95 5.2.3 AtBAG4 and TlBAG4 .................................................................................................... 95 5.2.4 Preparation of bacterial glycerol stock ........................................................................... 96 5.2.5 Extracting plasmid DNA (Mini prep) ............................................................................. 96 5.2.6 Plasmid TOPO

® Cloning ................................................................................................ 97

5.2.7 Transformation of bacteria.............................................................................................. 99 5.2.8 Restriction enzyme digestion of plasmid DNA .............................................................. 99 5.2.9 Agrobacterium-infiltration of Nicotiana tabacum and N. benthamiana ....................... 100 5.2.10 Histochemical GUS assay ............................................................................................. 100 5.2.11 Protein extraction .......................................................................................................... 100 5.2.12 SDS PAGE ................................................................................................................... 101 5.2.13 Coomassie blue staining ............................................................................................... 101 5.2.14 Western blotting ........................................................................................................... 102

vi Structural, physiological and molecular characterisation of the Australian native resurrection grass Tripogon


5.2.15 Protein microarray hybridisation (probing) and scanning ............................................ 102 5.2.16 Statistical analysis ........................................................................................................ 103 5.2.17 Bioinformics analysis ................................................................................................... 103 5.2.18 Quantitative real-time PCR analysis ............................................................................. 104

5.3 Results ..................................................................................................................................... 105 5.3.1 Generation of constructs for Agrobacterium-infiltration .............................................. 105 5.3.2 Agrobacterium transformation and infiltration of tobacco and N. benthamiana .......... 107 5.3.3 Protein extraction and purification ............................................................................... 109 5.3.4 Protein microarray hybridisation (probing) and scanning ............................................ 109 5.3.5 Statistical and bioinformatics analysis .......................................................................... 113 5.3.6 Generation of constructs for Agrobacterium-infiltration .............................................. 116

5.4 Discussion ................................................................................................................................ 118

CHAPTER 6: GENERAL DISCUSSION ...................................................................................... 125

6.1 Introduction.............................................................................................................................. 125

6.2 Structural and physiological features and changes of T. loliiformis leaves are to minimise

mechanical and oxidative stress dehydration and rehydration ............................................................ 125 6.2.1 Bulliform cells appear to play a significant role in gas exchange and leaf folding

during dehydration ........................................................................................................ 126

6.3 Tripogon loliiformis maintains its cellular integrity during dehydration and rehydration ....... 128 6.3.1 Increase electrolyte leakage is perhaps due to breakdown of macromolecules as

the result of autophagy during dehydration .................................................................. 129 6.3.2 Reduction of photosynthesis during early stages of dehydration plays a key role

in activation of autophagy procedure and increase in EL ............................................. 130 6.3.3 Plasma membrane is protected from mechanical damage during dehydration ............. 130 6.3.4 The possible protection of mitochondria structure during dehydration and

rehydration .................................................................................................................... 133

6.4 Protein-protein interaction profile of BAG4 suggest a key role in autophagy ......................... 134

6.5 Summary .................................................................................................................................. 136

REFERENCES .................................................................................................................................. 138

APPENDICES ................................................................................................................................... 156 Appendix A: Enriched GO terms for AtBAG4 and TlBAG4 .................................................. 156


loliiformis (F.Muell.) C.E.Hubb. during dehydration and rehydration vii

List of Figures

Figure 1: The classification of the land plants (embryophytes) based on water deficit

tolerance and classification of desiccation-tolerant plants. ............................................... 5

Figure 2: A schematic structure of C4 (Kranz) type XyMS+. ......................................................... 32

Figure 3: A schematic demonstration of enzyme compartmentalisation and structural

characteristics in C4 biochemical NAD-ME type photosynthesis. ................................. 32

Figure 4: Distribution of Tripogon loliiformis in Australia (Orchard and Wilson, 2005). ............ 33

Figure 5: The top panel shows Tripogon loliiformis in its natural habitat while the bottom

panel shows this plant grown from seeds in glasshouse in in 65mm pots. ..................... 35

Figure 6: Freehand transverse sections of hydrated and dehydrated leaves of Tripogon

loliiformis leaves.................................................................................................................. 60

Figure 7: Environmental scanning electron microscopy (ESEM) images of representative

hydrated leaves of Tripogon loliiformis. ............................................................................ 61

Figure 8: Difference between the process of leaf folding in middle and base of fully grown

Tripogon loliiformis leaves using environmental scanning electron microscopy

(ESEM). ............................................................................................................................... 65

Figure 9: Differences in internal and external structures from apical to basal regions of

Tripogon loliiformis leaves. ................................................................................................ 66

Figure 10: Changes in internal structure and chlorophyll content during dehydration and

rehydration of Tripogon loliiformis leaves. ....................................................................... 67

Figure 11: Changes in water content during dehydration and rehydration in Tripogon

loliiformis leaves.................................................................................................................. 78

Figure 12: Changes in chlorophyll fluorescence during dehydration and rehydration in

Tripogon loliiformis leaves. ................................................................................................ 78

Figure 13: Demonstrating cell membrane integrity in leaves using conductivity

measurement during dehydration and rehydration in Tripogon loliiformis

leaves. .................................................................................................................................. 81

Figure 14: Confocal laser-scanning microscopy of propidium iodide (PI) stained

hydrated, dehydrated (air-dry), rehydrated and control leaves of Tripogon

loliiformis. ............................................................................................................................ 82

Figure 15: Changes in photosynthetic rate during dehydration and rehydration in

Tripogon loliiformis. ........................................................................................................... 84

Figure 16: The experimental design of molecular work procedure. ............................................... 94

Figure 17: The position of the gene in pYL436 vector after cloning. .............................................. 98

Figure 18: Restriction enzyme digestion of the pENTR and pYL436 for verification of the

insertion of the constructs. ............................................................................................... 106

Figure 19: Determination of the optimal Agrobacterium-transient expression system. .............. 108

Figure 20: Peptide construct before and after isolation. ................................................................ 110

Figure 21: Verification of expression, extraction and isolation of target proteins using

Coomassie blue staining and Western blotting. ............................................................. 111

Figure 22: The procedure of protein microarray chip hybridisation. .......................................... 112

Figure 23: The scanned Arabidopsis thaliana protein microarray slides hybridised with

AtBAG4 and TlBAG4 constructs. ................................................................................... 114

viii Structural, physiological and molecular characterisation of the Australian native resurrection grass Tripogon


Figure 24: qRT- PCR results of TlBAG4 during dehydration and rehydration in Tripogon

loliiformis........................................................................................................................... 117

Figure 25: Comparison between human BAG4 BAG domain (hBAG4 BD) and

Arabidopsis thaliana BAG4 BAG domain (AtBAG4 BD). ............................................. 120

Figure 26: A schematic illustration of protein-protein interaction of AtBAG4 based on

experimental proven and bioinformatics prediction data based on string-

db.org. ............................................................................................................................... 121


loliiformis (F.Muell.) C.E.Hubb. during dehydration and rehydration ix

List of Tables

Table 1: The best studied angiosperm resurrection plants [modified from Bartels and

Hussain (2011)]: .................................................................................................................... 9

Table 2: The occurrence of resurrection plants in various genera and families among

Australian angiosperms (Gaff, 1981). ............................................................................... 29

Table 3: Abbreviations and location/section of Tripogon loliiformis leaf structures ..................... 58

Table 4: The density of stomatal complex, gland and prickle hair on adaxial and abaxial

surface of the leaf per mm2. ............................................................................................... 62

Table 5: The sequences of primers used for amplification of AtBAG4, TlBAG4 and

forward and reverse universal M13 primers used for sequencing. ................................ 96

Table 6: The comparison between the number of spots with significant signal values

between AtBAG4 and TlBAG4 hybridised Arabidopsis thaliana protein results. ....... 115

Table 7: The pathway enrichment analysis of proteins with significant signal values from

AtBAG4 and TlBAG4 microarray protein results. ....................................................... 115

x Structural, physiological and molecular characterisation of the Australian native resurrection grass Tripogon


List of Abbreviations

aa= amino acid(s)

APX = ascorbate peroxidase

ATP = adenosine-5’-triphosphate

bp = basepair(s)

CaMV = Cauliflower mosaic virus

cDNA = complementary DNA

CMA = Chaperone-mediated autophagy

CTCB = Centre for Tropical Crops and Biocommodities

dH2O = distilled water

DMSO = dimethyl sulfoxide

DNA = deoxyribonucleic acid

DT= desiccation tolerant

DTT = 1,4-dithiothreitol

DW = dry weight

EDTA = ethylenediaminetetraacetic acid

E. coli = Escherichia coli

EL = electrolyte leakage

ESEM= Environmental scanning electron microscopy

FAO = Food and Agriculture Organization

Fv/Fm = maximum photochemical quantum yield of Photosystem II

FW/DW = fresh-dry weight ratio

GFP = green fluorescent protein

GPS= global position satellite

LB = Luria-Bertani

mRNA = messenger RNA

MPa= megapascal

MS = Murashige & Skoog

NAD-ME= nicotinamide adenine dinucleotide-malic enzyme

N-terminal = amino-terminal

PBS = phosphate buffered saline

PCR = polymerase chain reaction


loliiformis (F.Muell.) C.E.Hubb. during dehydration and rehydration xi

pH = log (proton concentration)

PI = propidium iodide

qRT-PCR = quantitative reverse transcription polymerase chain reaction

QUT = Queensland University of Technology

RPM = rounds per minute

RNA = ribonucleic acid

RNase = ribonuclease

ROS = reactive oxygen species

Rubisco = ribulose-1,5-bisphosphate carboxylase/oxygenase

RWC = relative water content

SDS = sodium dodecyl sulphate

SOD= superoxide dismutase

TE = Tris-EDTA

TEMED = Tetramethylethylenediamine

TW = turgid weight

Tween 20 = polyoxyethylene (20) sorbitan monolaurate

UV = ultra violet

WT = wild type

Plant structures:

ew = Epicuticular wax

bu = Bulliform cell

g = Gland

lo = Long cell

pa = Papilla

pr = Prickle hair

pr-bc = Prickle-hair basal cell

si-s = Saddle silica-cell

st = Stomatal complex

sh-n = Nodular short-cell

bs = Bundle sheath

m = Mesophyll

mv = Metaxyleme vessel

xii Structural, physiological and molecular characterisation of the Australian native resurrection grass Tripogon


s = Sclerenchyma

p = Phloem

x = Xylem

XyMS+= Xylem mestome present

Units:

cm = centimetre(s)

⁰C = degree Celsius

d = day(s)

kDa = kilo Dalton(s)

g = gram(s)

h/hrs = hour(s)

l = litre(s)

M = molar

m = meter(s)

MW = molecular weight

min = minute(s)

mol = mole(s)

s = second(s)

V = volt(s)

vol = volume(s)

v/v = volume per volume

W = watt

w/v = weight per volume

Prefixes:

K = kilo

m = milli

μ = micro

n = nano


loliiformis (F.Muell.) C.E.Hubb. during dehydration and rehydration xiii

xiv Structural, physiological and molecular characterisation of the Australian native resurrection grass Tripogon


Statement of Original Authorship

The work contained in this thesis has not been previously submitted to meet

requirements for an award at this or any other higher education institution. To the

best of my knowledge and belief, the thesis contains no material previously

published or written by another person except where due reference is made.

Signature:

Date: _________________________

QUT Verified Signature


loliiformis (F.Muell.) C.E.Hubb. during dehydration and rehydration xv

Acknowledgements

I wish to thank my supervisory team Professor Sagadevan G. Mundree, Dr

Tanya Scharaschkin and Dr Brett Williams who are not only fantastic supervisors but

also I consider them my family. There are no words that I can sufficiently use to

express my gratitude towards them. They changed me for ever. Sagadevan, you are a

special person from whom I have learnt a lot, you are a role model in many aspects

of my life. Tanya, it was from you that I learnt scientific thinking and discipline. You

helped me to start my scientific journey. Brett, a true blue mate. You are like a

brother to me and I always admire your intelligence, gentle, down-to-earth

personality and no ego.

My special thanks go to Professor Emeritus Acram Taji who was like an angel

in my life, coming up with a miracle each time and saving me whenever I was in

trouble. I remember I was reading your books in Iran and asking myself would it be

possible if I would ever meet you… who would have thought that you would become

the reason I undertook my PhD.

There are a number of the people who were my closest friend who were there

whenever I needed help and with them I was most comfortable, Grace, Isaac, Karma,

Melody, My Linh, Purnika, Thita, Trang and the rest of M5 gang, H1 friends and

abiotic friendly stress group . I hope that I will always have such good friends near

me, friends are relatives that you choose.

There are so many people in the lab that I would like to thank. Hao who was

my big brother in the lab, Maiko a friendly protein advisor, Dani the queen of the lab

and so many other colleagues who helped me whenever I was lost in the lab.

Finally and most of all, I would like to thank my parents who because of them I

was able to carry on with this course and complete this thesis.

Thank you all!

xvi Structural, physiological and molecular characterisation of the Australian native resurrection grass Tripogon


Chapter 1: Introduction and Literature Review 1

Chapter 1: Introduction and Literature

Review

1.1 INTRODUCTION

At any given time, in any given year, there is always a part of the world that

is drought-declared. Combined with the additional pressure of feeding an ever

increasing global population that is expected to exceed nine billion by 2050

(Maxwell and Johnson, 2000), food security has become arguably one of the most

talked-about global concerns. More than one-third of the land in the world is located

in arid and semi-arid climates and in many countries, such as Australia, drought is

the major limitation for crop production (Gaff, 1981;Ali and Talukder, 2008). One

way to enhance food production during drought is to develop resilient crops. Genetic

engineering has emerged as a possible tool to produce drought-tolerant crops by

incorporating genes from plants which are able to tolerate drought into otherwise

susceptible crop plants (Ewen and Pusztai, 1999).

While most terrestrial plants are vulnerable to dehydration, certain plants

called “resurrection plants” can withstand periods of water deficit. The term

“resurrection plant” refers to those plants that use complex mechanisms that enable

them to lose over 85 % of their water and remain dormant until receiving water

(Gaff, 1971). There has been much interest in resurrection plants because they have

the potential to provide insights into drought-tolerance mechanism which has the

potential to be applied to agricultural crops.

Although molecular, physiological and to a lesser extent, structural responses

of resurrection plants to dehydration have been studied since 1912 (Irmscher), most

research has been limited to a small number of species, and these have been mainly

from southern Africa. Studying the resurrection behaviour of other species can

provide scientists with a new window of opportunity to understand this unique trait.

This PhD project examined one such species, T. loliiformis (F.Muell.) C.E.Hubb, that

is indigenous to Australia.

2 Chapter 1: Introduction and Literature Review

Tripogon loliiformis (family Poaceae) is an annual to short-lived perennial

resurrection grass (Gaff, 1981;Peterson et al., 2010). There has been very little

research carried out on this resurrection plant and our knowledge is limited mostly to

some ecological and general physiological studies (Gaff and Latz, 1978;Gaff and

McGregor, 1979;Gaff, 1981). Tripogon loliiformis possesses remarkable

characteristics that make it an ideal candidate as an experimental model for studying

the dehydration and rehydration responses (discussed at 1.8).

The aim of this study was to investigate the anatomical and physiological

response mechanisms of T. loliiformis during various stages of dehydration and

rehydration. Furthermore, a T. loliiformis orthologue of the well characterised anti-

apoptosis protein BAG4 (AtBAG4) from Arabidopsis thaliana was used in a protein

microarray for a comparative study with BAG4 from T. loliiformis (TlBAG4) in

order to understand the molecular interaction profile of this protein and to compare

the difference between TlBAG4 and AtBAG4. The outcomes of this project could

help provide further insight into abiotic stress tolerance and desiccation-tolerance

mechanisms. These outcomes also will be used as a platform for future studies on

this plant and could possibly be used to develop more tolerant crops.

1.2 CLASSIFICATION OF LAND PLANTS BASED ON THEIR

TOLERANCE TOWARD WATER DEFICIT

Most plants can be categorised into one of two groups: drought-sensitive plants

and drought-tolerant plants. Drought-sensitive plants, which include the majority of

agricultural crops, are vulnerable to even mild water deficit. Drought-tolerant plants

(also known as homioihydrous plants) can tolerate moderate levels of water deficit.

These plants keep their hydration level stable mainly through dehydration avoidance

mechanisms or pumping water from the soil via their deep root systems. However,

there is a third category, known as desiccation-tolerant plants, comprising plants that

can withstand extreme dehydration or desiccation and can lose over 85 % of their

relative water content RWC (Gaff, 1981;Hoekstra et al., 2001;Vicré et al., 2004a).

Although these plants are not of any significant agricultural importance, a better

understanding of the desiccation-tolerance mechanisms could be used to develop

drought-tolerant crops to meet the increasing global food demand.


1.3 DESICCATION TOLERANCE IN PLANTS

Desiccation-tolerance is defined as the ability of losing water to air-dry levels

and return to normal function after receiving water (Gaff, 1971). Although the

majority of plants are able to produce desiccation-tolerant seeds or spores, the ability

of tolerating desiccation in vegetative tissues is very rare (Bernacchia and Furini,

2004). Only a small portion of vascular plants and some non-vascular plants are able

to tolerate extreme desiccation and return to their normal metabolic function after

rehydration (Scott, 2000). These plants possess mechanisms and structures that

enable them to protect themselves against damage caused by extreme desiccation.

1.3.1 Evolutionary aspects

Desiccation-tolerance of vegetative tissues is thought to have evolved early in

the history of terrestrial plants as part of their adaptation to life on land (Oliver et al.,

2000a). While desiccation-tolerance remained common among prokaryotes (Potts,

1994) and nonvascular plants, this trait was subsequently lost possibly due to the

high energetic cost and/or the development of complex structures, such as the

evolution of vascular tissue, cuticle and stomata (Rascio and La Rocca, 2005).

However the genes related to desiccation-tolerance were maintained and these genes

are active in almost all land plants during reproduction. In vascular plants

desiccation-tolerance is mainly restricted to reproductive tissues such as spores,

pollen, orthodox seeds and some bulbs (e.g. Ranunculus bulbs). Desiccation-

tolerance in vegetative tissues appears to have ‘re-evolved’ independently, in

vascular plants, Selaginella, ferns, and at least eight times in angiosperms (Oliver et

al., 2000a;Proctor and Tuba, 2002).

1.3.2 Geographic distribution and ecology

Desiccation-tolerant plants are mostly small in size and slow-growing. They often

possess some features that are also exhibited by drought-tolerant plants (such as thick

cuticle, hairs or scales on leaves) (Gaff, 1981;Proctor and Tuba, 2002). Due to their

slow growth, these plants tend to grow where their resurrection capability is

optimally adaptive (Gaff, 1981). Desiccation-tolerant plants can mostly be found in

habitats where poor conditions, such as dryness, sporadic rain or shallow soil, limit

the growth of other plants (Iturriaga et al., 2000). In particular, on harsh

microhabitats of granitic and gneissic outcrops (inselbergs) of tropical and


subtropical regions, desiccation-tolerant vascular plants may become dominant

(Porembski and Barthlott, 2000;Bartels, 2005;Porembski, 2011). Most of the known

vascular desiccation-tolerant plants are from the southern hemisphere particularly,

from central and southern Africa and south-west Australia (Gaff, 1971; 1977;Gaff

and Latz, 1978). Desiccation-tolerant plants are classified in two groups: fully

desiccation-tolerant plants and modified desiccation-tolerant plants (Oliver and

Bewley, 1997) (Figure 1).


Figure 1: The classification of the land plants (embryophytes) based on water

deficit tolerance and classification of desiccation-tolerant plants.

Land plants can be divided into three main categories, i) drought-sensitive, ii) drought-

tolerant and iii) desiccation-tolerant. Desiccation-tolerant plants can be divided into

two categories of fully desiccation-tolerant plants and modified desiccation-tolerant

plants. Desiccation- tolerant plants can be further classified into two main categories,

poikilochlorophyllous (photosynthetic apparatus is lost during desiccation), and

homoiochlorophyllous (photosynthetic apparatus maintain during desiccation).

Land Plants (Embryophytes)

Drought sensitive Drought-tolerant Desiccation-tolerant

Homoiochlorophyllous

(Photosynthetic apparatus maintained)

Modified Desiccation-Tolerant Plants (Resurrection plants)

(Vascular)

Poikilochlorophyllous (Photosynthetic apparatus lost)

e.g. Tripogon loliiformis

Fully Desiccation-Tolerant Plants

(Non-vascular)

e.g. Xerophyta scabrida

e.g. Tortula ruralis


1.3.3 Desiccation-tolerant plants types

Fully desiccation-tolerant plants (poikilohydrous plants)

The ability of withstanding extreme desiccation is common among non-

vascular plants (Porembski and Barthlott, 2000). All the non-vascular desiccation-

tolerant plants are also called fully desiccation-tolerant plants. Fully desiccation-

tolerant plants have developed mechanisms to withstand very high rates of

dehydration but because they do not possess vascular tissue, their tolerance strategies

are different from those of vascular plants (Rascio and La Rocca, 2005). These plants

can dehydrate in a few minutes and remain in a dehydrated-dormant for up to 19

years (Alpert, 2000). They can also rehydrate quickly within minutes, and resume

normal function within a few hours (Alpert, 2000). Fully desiccation-tolerant plants

are mostly slow growing, possibly due to their constant readiness for any dehydration

which requires significant amounts of energy (Oliver, 2008). The small cell volume

and thick cell walls help to reduce physical stress during dehydration and rehydration

(Proctor and Tuba, 2002; Rascio and La Rocca, 2005). Fully desiccation-tolerant

plants are found from tropical to polar regions, on surfaces of rocks, shallow soils

and bark of the trees (Proctor and Tuba, 2002). Perhaps the most studied fully

desiccation-tolerant plant is Tortula ruralis which is a moss native to a wide range of

habitats (Schonbeck and Bewley, 1981;Oliver, 1991;Oliver et al., 1993;Tuba et al.,

1996a;Oliver et al., 2000b).

Modified desiccation-tolerant plants (resurrection plants)

All the vascular plants capable of tolerating extreme dehydration (below 13

% RWC) in their vegetative tissues are called modified desiccation-tolerant plants

(Gaff, 1971; 1977;Rascio and La Rocca, 2005;Toldi et al., 2009) or ‘resurrection

plants’ (personal communication with Professor Melvin Oliver). Modified

desiccation-tolerant plants tend to dehydrate and rehydrate more slowly than fully

desiccation-tolerant plants. If the time needed for inducing desiccation-tolerance is

not provided during dehydration, resurrection will not occur (Oliver, 1996). They can

tolerate less extreme desiccation and for a shorter duration (up to five years)

compared with fully desiccation-tolerant plants (Gaff, 1981;Alpert, 2000;Scott,

2000;Mundree et al., 2002).


Desiccation-tolerance in vegetative tissues of vascular plants has been found in

around one thousand ferns/fern allies and among some 400 angiosperms making up

less than 0.2 % of the total flora (Proctor and Pence, 2002;Rascio and La Rocca,

2005;Toldi et al., 2009;Bartels and Hussain, 2011;Porembski, 2011). Among

angiosperms, resurrection plants have been identified within both monocotyledonous

and dicotyledonous species while the majority are monocotyledons with few records

among the dicotyledons or other angiosperms (Gaff and Latz, 1978;Toldi et al.,

2009). Desiccation-tolerance within dicotyledons appears mainly in

Scrophulariaceae, Gesneriaceae and Myrothamnaceae families. Most resurrection

plants are herbaceous plants and the largest reported resurrection plant has been

Myrothamnus flabellifolia which is a small woody shrub (between 0.5 m and 1.5 m

tall) (Sherwin et al., 1998;Moore et al., 2007). No resurrection plant has been

reported among Gymnosperms and trees (Alpert, 2000). Desiccation-tolerance is

suggested to be a size limited trait as no resurrection plants has been reported over a

certain height (Bewley, 1982).

Among monocotyledonous plants, the resurrection trait appears in a wide range

of families that are not closely related to each other. Sometimes only one species in a

genus may be desiccation-tolerant (Gaff, 1981). Resurrection plants are mostly found

in arid and semi-arid areas of tropical and subtropical regions of the world,

particularly in Africa and Australia (Porembski and Barthlott, 2000;Rascio and La

Rocca, 2005;Toldi et al., 2009). However, they can also be found in humid forests

and temperate regions such as the resurrection grass, T. loliiformis that occurs in

tropical regions of Australia (Proctor and Tuba, 2002).

The following list includes the genera with resurrection species (number of

currently known desiccation-tolerant members in parentheses) based on Proctor and

Tuba (2002) with an update using Bartels and Hussain (2011) and Table 1

demonstrates the best studied angiosperm resurrection plants:


Angiosperms. Monocotyledons. Cyperaceae. Afrotrilepis (1), Carex (1),

Coleochloa (2), Cyperus (1), Fimbristylis (2), Kyllingia (1), Mariscus (1),

Microdracoides (1), Trilepis (1). Liliaceae (Anthericaceae). Borya (3). Poaceae.

Brachyachne (1), Eragrostiella (3), Eragrostis (4), Micraira (5), Microchloa (3),

Oropetium (3), Poa (1), Sporobolus (7), Tripogon (10). Velloziaceae. Aylthonia (1),

Barbacenia (4), Barbaceniopsis (2), Nanuza (1), Pleurostima (1), Vellozia (c. 124),

Xerophyta (c. 28). Dicotyledons. Myrothamnaceae. Myrothamnus (2). Cactaceae.

Blossfeldia (1). Acanthaceae. Talbotia (1), Gesneriaceae. Boea (1), Haberlea (1),

Ramonda (3), Streptocarpus (20). Scrophulariaceae. Chamaegigas (2),

Craterostigma (12), Ilysanthes (1), Limosella (1), Lindernia (< 30). Lamiaceae.

Micromeria (1), Satureja (1).


Table 1: The best studied angiosperm resurrection plants [modified from

Bartels and Hussain (2011)]:

Species Family Class Origin Poikilo/Homoi

Xerophyta viscosa Velloziaceae Monocot Southern Africa P

X. humilis Velloziaceae Monocot Southern Africa P

Mysothamnus

flabellifolia

Myrothamnaceae Dicot Southern Africa H

Sporobolus

stapfianus

Poaceae Monocot Southern Africa H

Eragrostis

nindensis

Poaceae Monocot Southern Africa P

Craterostigma

plantagineum

Scrophulariaceae Dicot Southern Africa H

C. wilmsii Scrophulariaceae Dicot Southern Africa H

Lindernia

brevidens

Linderniaceae Dicot East Africa H

Boea hygrometrica Gesneriaceae Dicot Africa, Asia

Australia

H

Haberlea

rhodopensis

Gesneriaceae Dicot Europe H


Classification of resurrection plants

Modified desiccation-tolerant plants can be divided into two groups depending

on whether or not they dismantle their photosynthetic apparatus during the

desiccation process. Those plants that retain their photosynthetic apparatus intact

during desiccation such as Sporobolus stapfianus and S. festivus are called

homoiochlorophyllous (Dingkuhn et al., 1999). Those that dismantle their

photosynthetic apparatus are called poikilochlorophyllous. All the resurrection

species of genera Eragrostis, Coleochloa and Borya are poikilochlorophyllous (Gaff,

1971;Proctor and Tuba, 2002) (Figure 1). The latter have to resynthesise their

chlorophyll following rehydration therefore it takes longer to recover their

photosynthetic activity (Müller, 2008). Poikilochlorophyllous plants can only be

found among monocots (Proctor and Tuba, 2002) (Figure 1). Some genera contain

homoiochlorophyllous and poikilochlorophyllous members, for example while S.

stapfianus and S. festivus retain considerable chlorophyll, S. pellucidus and S.

lampranthus do not retain chlorophyll during desiccation state (Gaff and Ellis,

1974;Gaff and Latz, 1978) providing a valuable ground for comparative study

between these two classes.


1.4 IMPACTS OF WATER DEFICIT ON PLANTS

Water deficit results in dehydration of the cells and this causes ion toxicity and

oxidative stress, and mechanical damage to the cell. Water loss leads to an increase

in concentration of free radicals and peroxides such as Clˉ, NO3ˉ, OHˉ and H2O2 in

the protoplasm. High concentrations of these components cause ion toxicity and

oxidative stress that can disrupt the cellular metabolism resulting in damaged

proteins, lipids, and DNA (explained in 1.6.1) (Mundree et al., 2002;Vicré et al.,

2004b;Rascio and La Rocca, 2005). Mechanical damage such as fusion of cell

membrane and loss of membrane integrity can be caused by severe dehydration

(Hoekstra et al., 2001).

1.4.1 Plant responses to water deficit

Non-desiccation-tolerant plants react to water deficit using mechanisms to

avoid dehydration or to reduce the damage caused by dehydration. These

mechanisms enable non-desiccation-tolerant plants to withstand moderate

dehydration (losses up to 60 % RWC of hydrated tissue) (Gaff, 1981). Desiccation-

tolerant plants have many common mechanisms with non-desiccation-tolerant plants

to limit damage during moderate dehydration (Gaff, 1981). After this stage DT plants

implement additional survival strategies to allow equilibration of their water content

to the level of the humidity of their surrounding environment (Hoekstra et al.,

2001;Vicré et al., 2004b;Toldi et al., 2009). These mechanisms allow desiccation of

vegetative tissues to air-dry levels and a seed-like state of anhydrases (life without

water) until more favourable conditions are encountered (Mundree et al.,

2002;Proctor and Tuba, 2002;Rascio and La Rocca, 2005).

While the majority of response mechanisms to moderate dehydration (above 60

% RWC) in desiccation-tolerant plants also occur in non-desiccation-tolerant plants,

desiccation-tolerant plants possess additional mechanisms to withstand severe

dehydration. The mechanisms of dehydration tolerance can also be classified into the

three groups: 1) structural responses, 2) physiological responses, 3) and molecular

responses.

Modified desiccation-tolerant plants and desiccation-sensitive plants possess

structural characteristics to avoid dehydration during environmental humidity

fluctuation. These characteristics can be seen regardless of environmental conditions.


For example, many plants store water in different parts, such as leaves, roots, tubers,

rhizomes or trunk. In some plants the rate of water loss is minimised through the leaf

surface, for example, by having leathery or needle-like leaves or leaves with thick

cuticle, hairs or scales. When water deficit is severe or lasts for an extended period,

these adaptations cannot prevent the plant from dehydration.

1.4.2 Water deficit response characteristics common among all plants

Moderate dehydration triggers a range of responses in plants. These responses

generally maintain the cell water content to avoid damage caused by dehydration and

protection against oxidative stress. These responses could be classified into three

main categories: 1) structural responses, 2) physiological responses, 3) and molecular

responses.

1.5 STRUCTURAL ASPECTS

Water deficit induces a number of structural changes in plants in order to

increase water use efficiency. The key changes include a reduction in canopy size

and root proliferation, increased cuticle thickness and trichome density as well as

reducing leaf surface area. Water uptake can be enhanced through root system

expansion and reducing the evapo-transpirational loss through reducing the canopy.

Reduction in canopy size can occur through reduction in the number and/or size of

the leaves and reduction in height of the plant loss which results in increasing

root/leaf ratio (Nagarajan and Nagarajan, 2010).

1.5.1 Leaf surface structures

An increase in the trichomes (such as macro-hairs) reduces the air-movement

over the surface as well as deflecting light, thus reducing surface evaporation, while

increased cuticle thickness is an established dehydration avoidance mechanism in

plants (Eglinton and Hamilton, 1967). Increased trichome number and cuticle

thickness may facilitate desiccation tolerance in resurrection plants by slowing down

the rate of dehydration, thus providing crucial time for implementation of desiccation

tolerance mechanisms. Furthermore, these structures may also aid tolerance by

mitigating light-induced damage of desiccated tissue.


Farrant et al. (2009) demonstrated that scale-less leaves of the fern Mohria

caffrorum during rainy season do not survive desiccation while scaled leaves that are

present during the dry season are desiccation tolerant. This study suggests that the

role of scales in desiccation-tolerance is through reducing dehydration rate and

masking the chlorophyll during dehydration and in the desiccated state. While there

are potentially other factors also involved in desiccation-tolerance in the leaves of M.

caffrorum during the dry season, the association of scales with desiccation tolerance

suggests their importance in desiccation-tolerance.

1.5.2 Reducing leaf surface area

While the above structural changes take a relatively long period of time, the

reduction of leaf surface area, a major morphological change that occurs during

environmental stress, happens in a short period of time. Reducing leaf surface area

occurs mostly through leaf folding and rolling (particularly among monocotyledons).

Leaf folding and rolling are two morphological changes which play a significant role

in stress-tolerance in higher plants (Kadioglu and Terzi, 2007;Kadioglu et al., 2012).

Leaf folding and rolling can be triggered by different biotic and abiotic stress factors.

Biotic stress factors such as viruses, bacteria, fungi and herbivores and abiotic stress

factors such as excessive radiation, heat, salts and dehydration (Reviewed by

Kadioglu et al., 2012).

Leaf rolling and folding could benefit the plant in two major ways during water

deficit. Firstly, it produces a micro-environment which increases the humidity close

to the leaf surface resulting in the reduction of transpiration rate (Clarke,

1986;Heckathorn and DeLucia, 1991;Tanimoto and Itoh, 2000). Secondly, leaf

folding and rolling reduces leaf surface area resulting in less exposure to the light.

This results in less heat (Schakel and Hall, 1979;Turner et al., 1986;Heckathorn and

DeLucia, 1991;Turgut and Kadioglu, 1998), radiation damage (O'Toole and Cruz,

1980;Corlett et al., 1994;Omarova et al., 1995;Turgut and Kadioglu, 1998), and

photosynthesis-induced oxidative stress (O'Toole and Cruz, 1980;Sarieva et al.,

2010).

Leaf rolling has been extensively studied in monocotyledons, particularly in

major true cereals including maize (Fernandez and Castrillo, 1999;Nelson et al.,

2002), rice (O'Toole and Cruz, 1980;Turner et al., 1986;Heckathorn and DeLucia,


1991;Ekanayake et al., 1993;Price et al., 1997;Dingkuhn et al., 1999;Tanimoto and

Itoh, 2000;Singh et al., 2011), wheat (Clarke, 1986;Omarova et al., 1995;Dingkuhn

et al., 1999;Sarieva et al., 2010), sorghum (Matthews et al., 1990;Corlett et al., 1994)

and barley (Turner and Stewart, 1986). Ctenanthe setosa has been used as a

monocotyledonous model plant for studying leaf rolling in a number of reports

(Turgut and Kadioglu, 1998;Kadioglu and Turgut, 1999;Terzi and Kadioglu,

2006;Nar et al., 2009). The majority of these reports, investigate the molecular and

physiological aspects and only a few look at the structural aspects of leaf rolling in

monocotyledons (Esau, 1965;Alvarez et al., 2008).

While leaf movement in eudicotyledons is known to be controlled mainly by

pulvini cells (Pedersen et al., 1993;Taya, 2003), in monocotyledons, leaf rolling is

believed to be controlled by bulliform cells in the presence or absence of colourless

cells which act as pivots. Loudetiopsis chrysothrix and Tristachya leiostachya are

prime examples of the latter phenomenon (Esau, 1965;Alvarez et al., 2008). Vecchia

et al. (1998) however argue that in S. stapfianus (Poaceae) dehydration of bulliform

cells does not result in any significant leaf folding or rolling. A possible correlation

between sclerenchymous tissue and leaf rolling has been suggested in two

independent molecular studies on mutant rice (Zhang et al., 2009) with defective

sclerenchyma cells and in mutant maize (Nelson et al., 2002) with different

sclerenchymous tissue distribution. Despite some progress, the exact role of

sclerenchymous tissue in leaf folding and rolling in monocotyledons is still not clear.

Reducing leaf surface area is particularly important in resurrection plants

(Gaff, 1981;Sherwin et al., 1998;Vander Willigen et al., 2003;Moore et al.,

2007;Georgieva et al., 2010). Reducing leaf surface area is a dehydration avoidance

mechanism which slows down the dehydration rate thus providing more time needed

for inducing desiccation-tolerance (Vecchia et al., 1998;Oliver et al., 2000a). Leaf

folding and rolling can also reduce the damage caused by excessive radiation during

desiccation. Furthermore, less light exposure reduces photosynthesis and lowers the

amount of reactive oxygen species (ROS) produced as a result of photosynthesis

during water deficit stress (Farrant, 2000;Farrant et al., 2003). Leaf folding has been

reported in a number of resurrection plants (Gaff, 1981;Sherwin et al., 1998;Vander

Willigen et al., 2003;Moore et al., 2007;Georgieva et al., 2010), however, the


mechanisms of leaf folding have not been well-studied in monocotyledons. Farrant et

al. (2003) demonstrated that the leaves of resurrection plant Craterostigma wilmsii

that were mechanically prevented from folding, could not survive desiccation.

1.5.3 Xylem tissue (in stem and root)

A mature xylem tissue is dead with rigid cell walls, therefore, during

desiccation these cells are dry and filled with air. Water refilling of the long air-filled

vessels seems to be a major challenge for the resurrection plants. This xylem refilling

problem might contribute to the height limitation of the resurrection plants

(Porembski and Barthlott, 2000;Proctor and Tuba, 2002). In a number of resurrection

plants it has been reported that the re-greening does not take place on the tip of the

leaves and stems. Sherwin et al. (1998) explains that the distal portion of some

branches of Myrothamnus abellifolius did not resurrect after ground watering,

however when these parts were cut and placed in water, rehydration took place

within 12 h. This explains that the lack of re-greening on these might be due to the

problem of water reaching the tip of the branches rather than tissue damage.

Myrothamnus abellifolius is the only resurrection plant with a woody stem. In

a study by Sherwin et al. (1998), it was revealed that root pressure was not sufficient

to refill the xylem (only 24 cm) of desiccated M. abellifolius; the capillary force

however, was enough to rise the water in the xylem tissue (to 2.12 m). This

unusually strong capillary force is due to very narrow xylem vessels (one of the

narrowest xylems among seed plants) (Sherwin et al., 1998).

There is little published data about the changes in roots during desiccation in

resurrection plants; however it seems that rehydration does not take place solely due

to root pressure. The rehydration of the leaves in resurrection plants has been

demonstrated to occur both by watering roots or directly to the leaves, even watering

of the excised leaves can lead to re-greening (Gaff and Latz, 1978). Furthermore, this

strengthens the idea that resurrection plants do not rely solely on root pressure for

rehydration. While the embolism of the xylem tissue is said to be an obstacle in

resurrection in some resurrection plants (Sherwin and Farrant, 1996), perhaps

capillary force combined with the root pressure makes the refilling of the xylem

tissue possible (Sherwin et al., 1998).


Protection of the vascular tissue during desiccation is another vital issue for the

survival of resurrection plants. No significant changes in the vascular tissue of M.

abellifolius during desiccation were observed (Sherwin et al., 1998), however

dehydration stress damaged the vascular tissue of barley (Pearce and Beckett, 1987).

Unusual knob-like structures that were observed on the outer surface of xylem

vessels of M. abellifolius are suggested to have a possible role in protection of xylem

vessels during dehydration (Sherwin et al., 1998).

1.6 PHYSIOLOGICAL ASPECTS

Plants display a variety of physiological responses at the cellular and whole-

organism levels during dehydration and rehydration. Some of the key changes

include photosynthesis, stomata closure, pigmentation and respiration. These changes

are mainly to prevent oxidative damage resulting from ROS accumulation during

dehydration.

1.6.1 Reactive oxygen species

Accumulation of ROS is one of the major problems during dehydration.

Reactive oxygen species such as superoxide, hydrogen peroxide and hydroxyl

radicals damage macromolecules in the protoplasm (Bowler et al., 1992;Smirnoff,

1993). Photosynthesis and respiration are the main sources of ROS during drying

resulting from the interruption of electron transport (Hendry and Grime,

1993;Smirnoff, 1993;Farrant, 2000;Mundree et al., 2002;Apel and Hirt, 2004b).

Reactive oxygen species are formed naturally as a byproduct of photosynthesis

and respiration and neutralised through antioxidants in the cell. During the stress

such as water deficit, however, the production of ROS exceeds the capacity of

antioxidant system (such as Mehler ascorbate peroxidase pathway), leading to rapid

increase in ROS concentration which is called “oxidative burst”. During water deficit

stress, the availability of the CO2 is restricted (stomata closure and/or low amount of

water), in this condition Rubisco favours oxygen over CO2 to pass the electron (Apel

and Hirt, 2004a). This leads to production of O2ˉ which can attack and produce other

reactive oxygen species such as Oˉ, OHˉ and H2O2. High concentrations of ROS can

cause ion toxicity and oxidative stress that can disrupt the metabolic procedure of the


cell and damage proteins, lipids, and DNA (Mundree et al., 2002;Vicré et al.,

2004b;Rascio and La Rocca, 2005). Reactive oxygen species also can activate the

apoptosis pathway and lead to tissue senescence.

In order to prevent ROS accumulation, resurrection plants stop photosynthesis

through different ways. The first step to reduce photosynthesis is through stomata

closure. Closed stomata limits the gas exchange and lowers CO2 level which in turn

reduces photosynthesis (Tuba et al., 1998). Lowering of the photosynthesis through

stomata closure is particularly effective in C3 plants. Furthermore, the closed stomata

reduces the water lose through transpiration. In a study by Moore et al., (2007) it was

shown that in Myrothamnus flabellifolia the stomata remained open irreversibly upon

desiccation for secretion of calcium salts. The steps step/s in reducing photosynthesis

in resurrection plants is through dismantlement of photosynthetic apparatus and/or

prevention of light-chlorophyll interaction through pigmentation and/or reducing leaf

surface area.

1.6.2 Photosynthesis

In comparison with mesophytes, resurrection plants respond to dehydration

faster and reduce photosynthesis relatively early from 80 % RWC and below

(Farrant, 2000). If dehydration continues, resurrection plants terminate

photosynthesis. In poikilochlorophyllous resurrection plants, chlorophyll pigments

are disintegrated and thylakoid membranes are dismantled to prevent the production

of ROS through photosynthesis. Homoiochlorophyllous resurrection plants retain

much of their photosynthetic apparatus but limit photosynthesis through shading,

pigmentation and/or reduction of leaf surface area (Hallam and Luff, 1980;Gaff,

1981;Sherwin and Farrant, 1998;Tuba et al., 1998;Farrant, 2000;Farrant et al., 2003).

Down-regulation of photosynthesis-related gene expression during early stages of

dehydration is also shown to be involved in limiting photosynthesis in the

resurrection plant Craterostigma plantagineum (Bernacchia et al., 1996;Bockel et al.,

1998).

Pigmentation of leaves during dehydration is very common among

homoiochlorophyllous resurrection plants such as in T. loliiformis, C. wilmsii and M.

flabellifolius (Gaff, 1981;Sherwin and Farrant, 1998;Farrant, 2000). Reducing leaf

surface area through leaf folding and rolling is also common among


homoiochlorophyllous resurrection plants. Farrant et al. (2003) demonstrated that the

leaves of homoiochlorophyllous resurrection plant C. wilmsii cannot resurrect if the

leaves were mechanically prevented from folding during desiccation as a result of

light-damage. The pigmentation on the exposed surfaces of the leaves and leaf

surface area prevent light-chlorophyll interaction and minimise light-induced

damage.

1.6.3 Respiration

Respiration has been shown to remain active until very low hydration levels in

some resurrection plants, perhaps for providing energy needed for protection

mechanisms when photosynthesis is shut down (Tuba et al., 1996b;Tuba et al.,

1997;Tuba et al., 1998;Farrant, 2000;Whittaker et al., 2004). Respiration has been

shown to be inhibited in the whole-plant under dehydration stress in desiccation

sensitive plants (Leprince et al., 2000). However, similar to resurrection plants

mitochondrial respiration remains active in seeds until very low hydration levels

(Benamar et al., 2003;Atkin and Macherel, 2009). Mitochondrial respiration resumes

very rapidly in seeds as well as in homoiochlorophyllous resurrection plants (Tuba et

al., 1998).

The quick resumption of mitochondrial respiration as well as a structural

observation by Farrant (2000) suggests that membrane integrity of mitochondria in

homoiochlorophyllous resurrection plants is preserved during dehydration.

Membrane integrity of the mitochondria has been shown to be protected during

desiccation in orthodox seeds (Benamar et al., 2003). While mitochondria in the

poikilochlorophyllous resurrection sedge Coleochloa setifera had fewer cristae

during desiccation (Bartley and Hallam, 1979), in four species of

poikilochlorophyllous resurrection plants from the Borya genus, mitochondria lost

some of the peripheral membrane and the number of cristae reduced (Gaff, 1981).

The resumption of respiration in poikilochlorophyllous resurrection plant X. scabrida

was shown by Tuba et al. (1998) after 24 h.

These studies suggest that homoiochlorophyllous resurrection plants maintain

their mitochondrial membrane integrity resulting in rapid resumption of respiration.

In contrast, poikilochlorophyllous resurrection plants lose at least a part of their

mitochondrial membrane integrity and resume the respiration with a delay. However


this area of study requires further research particularly on the resumption of

respiration in homoiochlorophyllous resurrection plants as the only work on

resumption of respiration in homoiochlorophyllous plants has been on a moss and

lichen and not on any vascular homoiochlorophyllous resurrection plants.

1.7 MOLECULAR RESPONSES TO WATER DEFICIT IN PLANTS

Molecular responses to moderate dehydration in plants play three main roles:

1) structural and osmoprotection, 2) oxidative and ion toxicity protection, 3) and

regulatory. During dehydration, cells produce various osmolytes to maintain water

content and to protect the structure of its components. Some of these molecules are

carbohydrates (sucrose in particular) and a range of hydrophilic proteins such as late

embryogenesis abundant (LEA) proteins and small heat shock proteins (HSPs)

(Almoguera et al., 1993;Rascio and La Rocca, 2005). Synthesis of antioxidants and

ion scavengers such as ascorbates, peroxidises and superoxide dismutases protect the

cell against ion toxicity and oxidative stress (Hara et al., 2004;Mhadhbi et al., 2011).

The molecular responses to water deficit are induced either directly by loss of

water or by regulatory signals (Rascio and La Rocca, 2005). Roots act as the primary

sensors to regulate plant responses (Tardieu, 1996). Abscisic acid (ABA) plays a

major regulatory role in vascular plants during water deficit (Bray, 2002;Osakabe et

al., 2014). This signal-molecule is produced in the roots and regulates numerous

molecular responses in the plant during water deficit such as, antioxidant production

and expression of most of the dehydration-related genes or triggers physiological

responses such as stomatal closure and root expansion (Guan et al., 2000;Rascio and

La Rocca, 2005;Geisler et al., 2006).

1.7.1 Molecular responses to water deficit in desiccation-tolerant plants

While molecular responses to dehydration in desiccation-tolerant plants have

many common components with desiccation sensitive plants, desiccation-tolerant

plants possess additional mechanisms to withstand severe dehydration. These

molecular responses in desiccation-tolerant plants in response to dehydration can be

roughly divided into four groups while some molecules can be in more than one

group; 1) regulatory molecules, 2) antioxidants, 3) carbohydrates and 4) proteins.


This section of the literature review discusses these four groups of molecular

responses in desiccation-tolerant plants.

1.7.2 Regulatory

During desiccation, regulation of responses is generally classified as ABA-dependent

and ABA-independent.

ABA-dependent signalling

Abscisic acid is the key signal molecule during severe dehydration just like

during moderate dehydration. ABA concentrations in leaf cells can increase by 3- to

20-fold upon desiccation (Gaff and Loveys, 1984;Dingkuhn et al., 1999;Bernacchia

and Furini, 2004;Dinakar et al., 2012). Several proteins have been identified to be

synthesised both by desiccation or induced by exogenous ABA-treatment in C.

plantagineum while the expression levels of these proteins declined after relief from

stress and ABA-treatment (Bartels et al., 1990;Piatkowski et al., 1990;Michel et al.,

1994). The desiccation sensitive callus of C. plantagineum turns desiccation-tolerant

when treated with exogenous ABA (Bartels et al., 1990). Similarly ABA is

accumulated in orthodox seeds and is involved in desiccation-tolerance and

dormancy of the seeds (del Carmen Rodríguez-Gacio et al., 2009). While the role of

ABA is well studied in desiccation-tolerance, the role of other plant hormones such

as jasmonic acid, ethylene, auxin and salicylic acid is not well understood. Most of

these hormones are related to senescence and breaking the dormancy which relieves

desiccation-tolerance, however they might have a role in resurrection and renewing

metabolism after receiving water.

ABA-independent signalling

Transcription factors control the responses at the cellular level by regulating

gene expression (Neale et al., 2000;Le et al., 2007). It is suggested that the re-

evolvement of desiccation-tolerance in the vegetative tissues of angiosperms is the

result of several genes which are active in dry reproductive tissues (e.g. seed and

pollen) (Oliver et al., 2000a). This might mean that specific transcriptional factors in

resurrection plants regulate the expression of these genes in the vegetative tissues. A

study using transcriptome analysis by next-generation sequencing revealed that

members of the NAC, NF-YA, MADS box, HSF, GRAS, and WRKY transcription


factor families play a key role in diverse physiological and molecular changes in

resurrection plants leading to desiccation-tolerance (Gechev et al., 2013). Phillips et

al. (2002) have found a new family of genes called Plastid Targeted Protein (CpPTP)

with a possible role in transcriptionally down-regulating the plastid genome during

desiccation by binding to DNA in C. plantagineum.

Small RNAs have shown to have an essential role in gene regulation for

environmental and developmental changes (Phillips et al., 2007). The role of small

RNAs in desiccation-tolerance have been revealed in C. plantagineum (Hilbricht et

al., 2008). It is expected more information about the role of small RNAs in

desiccation-tolerance be revealed in the near future. Some responses could be

independent of molecules and may be induced directly by dehydration such as the

expression of the XVSAP1 gene (Garwe et al., 2003). Even though XVSAP1 can also

be expressed through ABA signalling (Iyer et al., 2008).

1.7.3 Antioxidants

One of the main problems plants face during dehydration is the oxidative stress

due to accumulation of ROS. One of the key strategies for protecting the cell against

oxidative stress during the dehydration is by increasing the synthesis of antioxidants.

Enzymes such as catalases, superoxide dismutase (SOD), ascorbate peroxidase

(APX) and glutathione reductase act as ROS scavengers. Additionally, non-

enzymatic antioxidants such as ascorbate, glutathione, carotenoids, anthocyanins,

polyphenols, osmolytes, proteins (e.g. peroxiredoxin) and amphiphilic molecules

(e.g. tocopherol) also regulate ROS levels (Bowler et al., 1992;Navari-Izzo et al.,

1997;Noctor and Foyer, 1998;Sherwin et al., 1998;Sgherri et al., 2004). The

antioxidant activity needs to remain active until a low hydration level to protect the

cell from ROS produced by respiration (Sherwin and Farrant, 1998;Farrant, 2000).

Antioxidant activity must be particularly critical for homoiochlorophyllous

resurrection plants as the maintained chlorophyll could be a source of ROS

production during desiccation. Kranner et al. (2002) demonstrated an increased

accumulation of antioxidants in desiccated M. abellifolia. This study also showed the

amount of broken down antioxidant in the desiccated tissue has a direct relation with

the period of desiccation. In many homoiochlorophyllous resurrection plants leaves


are heavily pigmented mostly by anthocyanin (e.g. C. wilmsii and T. loliiformis) that

has antioxidative property and also protects the tissue from radiation from the sun

during desiccation (Gaff, 1981;Sherwin and Farrant, 1998). A novel member of the

vicinal oxygenase chelate (VOC) superfamily of metalloenzymes was found to

accumulate during both dehydration of vegetative tissues and dry seeds of X. humilis

(Mulako et al., 2008).

1.7.4 Carbohydrates level in desiccation tolerant plants in resurrection plants

Carbohydrate production is one of the main responses to water deficit in a wide

range of plants (Schakel and Hall, 1979). Sucrose content increases more than other

carbohydrates during water deficit stress (Schakel and Hall, 1979;Turner et al.,

1986). It has been suggested that carbohydrates have an osmoregulatory role during

minor water loss (Schakel and Hall, 1979). While carbohydrate production is one of

the major plant responses to dehydration, where accumulation of carbohydrate plays

a fundamental role in desiccation-tolerance in plants and other organisms. Non-

reducing oligosaccharides and sucrose in particular are the main carbohydrates

accumulated in seeds, pollen and in most of the plants that can tolerate desiccation.

In nearly all of the resurrection (vascular DT) plants studied to date as well as

other organisms that can withstand desiccation such as bacteria, fungi and yeast, the

levels of non-reducing disaccharides increase upon desiccation (Clarke, 1986;Crowe

et al., 1998;Nar et al., 2009;ElSayed et al., 2014). Carbohydrates play an important

part in structural protection during extreme dehydration. During extreme

dehydration, the viscosity of the cytoplasm increases dramatically (glassy state or

vitrified state), carbohydrate molecules form a protective layer around other

molecules and parts of the cell and act as a chaperone (Crowe et al., 1998). This

results in protection of these components from structural damage such as structural

damage to macromolecules, membrane fusion and molecular stability during severe

dehydration (Crowe et al., 1998;Bryant et al., 2001). A range of non-reducing

oligosaccharides (e.g. sucrose, sorbitol, trehalose and the raffinose family of

oligosaccharides) accumulate in resurrection plants mainly through breakdown of

starch during dehydration (Crowe et al., 1998;Mundree et al., 2000;Crowe et al.,

2001;Peters et al., 2007;ElSayed et al., 2014). Carbohydrate has also been shown to

protect membrane structure and preserving the fluidity of the plasma membrane in


dry seeds (Wang et al., 1999). Herein the role of some of these sugars in desiccation

tolerance is discussed.

Sucrose

Sucrose is the only cellular protection carbohydrate available in desiccated

tissues in fully desiccation tolerant mosses including Dicranum majus, Hookeria

lucens, Polytrichum commune, Racomitrium lanuginosum, Thuidium tamariscinum

and T. ruraliformis (O'Toole and Cruz, 1980). The sucrose level does not change

during dehydration and rehydration whether in the light or dark, while there is a

reduction in the amount of glucose and fructose (reducing carbohydrates) during

dehydration (O'Toole and Cruz, 1980). The sucrose content in the hydrated tissue of

desiccation tolerant mosses seems to be sufficient for protecting the cells during

desiccation (10 % of dry mass in T. ruralis). Furthermore, lack of change in its level

in desiccation appears to be a common feature in fully desiccated tolerant mosses

(O'Toole and Cruz, 1980). These characteristics could possibly explain the constant

readiness of facing desiccation in desiccation tolerant mosses. Furthermore, reducing

carbohydrates have shown to have browning and denaturising effects on proteins in

the anhydrous state (Turner and Stewart, 1986). Reduction in the amount of reducing

carbohydrates perhaps may help with reducing the chance of oxidative damage

during desiccation.

In modified desiccation-tolerant plants and also among all non-reducing

oligosaccharides, sucrose is the main product (Bianchi et al., 1991;Proctor and Tuba,

2002;Nar et al., 2009) and the predominant carbohydrate involved in mechanisms of

protection (Scott, 2000). However, there are exceptions such as a fern-ally

resurrection species Selaginella lepidophylla in which the main protectant

carbohydrate is trehalose (Alvarez et al., 2008). The concentration of sucrose can

increase up to 20 fold compared to hydrated state or 40 % of dry mass upon

desiccation (Dominy et al., 2008). Accumulation of sucrose is one of the last

preparations for cell protection for the desiccation period. This accumulation sucrose

starts below 20 % RWC in most of resurrection plants, while others start at below 60

% RWC (Toldi et al., 2009). This is when the cell is fully committed to a period of

dormancy and the concentration of sucrose coincides with the level of ABA.


In addition to the osmoregulatory role of sucrose in drying plants, it has been

suggested that sucrose can protect the integrity of the cell membrane by hydrogen

bonding to the polar head groups of phospholipid bilayers, and preventing damage or

fusion by replacing water and maintaining the spacing between them (Crowe et al.,

1998;Ghasempour et al., 1998). Suzuki et al. (1993) demonstrated that sucrose

stabilises enzymes to a great extent in the amorphous state. Sucrose (and other

carbohydrates accumulated during dehydration) could also be used as a source of

energy during rehydration and the recovery phase (Ghasempour et al., 1998).

Trehalose

Trehalose, is a rare non-reducing disaccharide found in desiccation tolerant

species from prokaryotes (Potts, 1994) to eukaryotes. Trehalose has been found to

accumulate in viable desiccated tissues of almost all studied modified desiccation-

tolerant species with occasional exceptions such as M. flabellifolius, X. villosa, S.

atrovirens and B. hygroscopica (Drennan et al., 1993;Iturriaga et al., 2000;Nar et al.,

2009). This carbohydrate is known to be one of the sources of energy and protection

in most living organisms and can be found in many organisms, including bacteria,

fungi (up to 10–25 % by dry weight in mushrooms), insects, invertebrates and

mammals however it is extremely rare in seed plants (Knapp, 1985;Crowe et al.,

1998;Hovakimyan et al., 2012). Certain crops transformed to produce trehalose have

shown tolerance to abiotic stressors that were associated with sustained plant growth

(Jang et al., 2003;Taya, 2003).

This non-reducing disaccharide has unique properties: it is the most stable

saccharide because of its high thermostability, a wide pH-stability range; does not

show Maillard reaction (non-enzymatic browning) with acids or proteins as it is a

non-reducing carbohydrate (Knapp, 1985); and anhydrous forms of trehalose readily

regain moisture to form the dehydrate. Trehalose also has diverse biological roles:

like sucrose, it is an effective protein stabiliser that preserves the integrity of

membranes during desiccation (Crowe et al., 1998). Trehalose is also a

transcriptional factor regulator (by attaching to protein's active site) and an allosteric

carbohydrate metabolism inhibitor (Zhang et al., 2009) it is also a regulator in a

various developmental stages particularly regulating metabolic responses during

stress (Paul et al., 2008;Gazzarrini and Tsai, 2014) including sucrose-starch balance

during dehydration (Lunn et al., 2014). Due to its chemically unreactive properties


trehalose has been suggested to accumulate to large concentrations without affecting

cellular metabolism (Toldi et al., 2009). These unique characteristics of trehalose

suggest it to be ideal as a protective carbohydrate during desiccation in resurrection

plants.

Raffinose

Raffinose family oligosaccharides (RFOs) have been reported to accumulate in

desiccated tissues of resurrection plants. They are good storage molecules and as

non-reducing carbohydrates they can accumulate in large quantities without

impacting critical metabolic processes (Peters et al., 2007). Stachyose is one of the

most important RFOs that contributes to 50 % of the carbohydrate in desiccated roots

of C. plantagineum (Bartels and Salamini, 2001;Norwood et al., 2003). Müller et al.

(1997) suggested the accumulation of raffinose in drying tissues of resurrection

plants might be a pre-adaptation strategy as raffinose can prevent sucrose

crystallisation. It has also been suggested that RFOs contribute in specific ROS

processes as well as disease prevention in plants (Van den Ende and Valluru, 2009).

The accumulation of non-reducing oligosaccharides seems to have a critical

role in desiccation-tolerance in different ways. Apart from the structural protection,

the non-reducing properties of these carbohydrates perhaps make them a chemically

stable carbohydrate during the oxidative stress (reducing) during dehydration.

Sucrose, trehalose and RFOs are known to have regulatory and ROS scavenging

roles in vascular plants (Paul et al., 2008;Van den Ende and Valluru, 2009;Gazzarrini

and Tsai, 2014), however to my knowledge these properties have not been

investigated in resurrection plants during drying.

1.7.5 Proline and proteins

Proline is a low-molecular-weight osmolytes which accumulates during water

deficit in plants. Accumulation of proline is involved with the osmotic adjustment in

the cell (Wahid and Close, 2007). In plants, calcium and auxin have been identified

in signalling mechanisms participating in water deficit-induced proline accumulation

(Farooq et al., 2009). Proline has been shown to increasingly accumulate during

severe dehydration (<40 % RWC and particularly <20 % RWC) and its concentration

reduced during rehydration in E. nindensis (Vander Willigen et al., 2004).


Furthermore, proline has been shown to accumulate in the small fragmented vacuoles

during desiccation and not the cytosol of the E. nindensis (Vander Willigen et al.,

2004). A proline-rich protein has also been shown to accumulate during desiccation

in S. stapfianus under severe dehydration (Neale et al., 2000). Perhaps proline is also

involved with the vitrification state and protection of the membrane and other

macromolecules in desiccation tolerant plants.

Protein production in desiccation sensitive plants ceases at mild levels of

dehydration. However in resurrection plants, protein production is essential during

dehydration and continues until the plant has reached an almost air-dry state (Bartels

et al., 1990;Gaff et al., 1997;Proctor and Pence, 2002). The soluble protein content of

S. stapfianus in desiccation-tolerant attached drying leaves doubles while it does not

change in detached desiccation sensitive drying leaves (Whittaker et al., 2004).

Furthermore, breakdown of the proteins for recycling the components for translation

of proteins involved in desiccation-tolerance seems to be essential in resurrection

plants (Griffiths et al., 2014).

While the majority of the proteins expressed in desiccation-tolerant plants are

similar to desiccation sensitive plants, proteomic comparison of S. stapfianus with a

closely related desiccation sensitive species, S. pyramidalis revealed a set of 12 novel

proteins that might be associated with desiccation-tolerance (Gaff et al., 1997).

Gechev et al. (2013) identified five transcriptional factors that are involved in

vegetative desiccation-tolerance. The main roles of these proteins expressed during

extreme dehydration in desiccation-tolerant plants can be categorised in at least one

of the following groups. 1) protein expression, 2) structural protection (chaperones),

3) osmotic adjustment (osmolytes) or 4) ROS scavenging. Here some of the key

proteins expressed during desiccation and their role in desiccation-tolerance are

discussed.

LEA proteins

Late-embryogenesis abundant (LEA) proteins (e.g. dehydrins) play a critical

role in establishing desiccation-tolerance in mature embryos (Close, 1996). They are

also expressed during osmotic stress as a response to ABA in plants and can also be

found in all other kingdoms (Proctor et al., 1998). High levels of LEA proteins in

viable desiccated tissues of plants as well as desiccation-tolerant animals have been


reported, showing the important role of this protein in desiccation-tolerance

(Tunnacliffe et al., 2005;Liu et al., 2009). Transgenic rice expressing a LEA protein

showed drought and salt-tolerance improvement (Xu et al., 1996;Chandra Babu et

al., 2004;Xiao et al., 2007). Late-embryogenesis abundant proteins are extremely

hydrophilic and resistant to denaturation (Proctor et al., 1998). They act as chaperone

or water hydration ‘shell’ around other molecules and protect them from denaturation

(Olvera-Carrillo et al., 2010). They can prevent unfavourable molecular interactions

or biochemical activities (Olvera-Carrillo et al., 2010).

HSPs

Small heat shock proteins (HSPs) accumulate to high quantities in desiccating

vegetative tissues in resurrection plants and believed to play a key role in acquiring

desiccation-tolerance (Alpert and Oliver, 2002;Gechev et al., 2013). These proteins

are also expressed during the final stages of the seed dehydration as well as in

response to heat stress in vegetative tissues (Singla et al., 1998;Kotak et al., 2007).

HSPs have shown to protect membrane and macromolecules from damaging effects

of desiccation (Sales et al., 2000). The protective ability of HSPs is believed to be

related to chaperone -like activities of these proteins (Alpert and Oliver, 2002).

Transgenic rice overexpression of HSP101 from Arabidopsis showed significant

improvement in growth at high temperature (Katiyar-Agarwal et al., 2003).

ELIPs

Early light-inducible proteins (ELIPs) have been observed to accumulate in

desiccated tissues of homoiochlorophyllous desiccation tolerant plants such as T.

ruralis, C. plantagineum, Haberlea rhodopensis and Selaginella tamariscina (Bartels

et al., 1992;Zeng et al., 2002;Liu et al., 2008;Gechev et al., 2012). ELIPs also

accumulate in plant tissues in response to various environmental stresses such as

high light, UV-B, methyl jasmonate, cold, low oxygen and CO2 concentration,

nutrient starvation, senescence (Zeng et al., 2002). ELIPs are mediated by ABA and

light (Bartels et al., 1992) and are thought to have a key role in protection/or

repairing of the photosystem (Ouvrard et al., 1996;Zeng et al., 2002). To my

knowledge only homoiochlorophyllous desiccation-tolerant plants accumulate ELIPs

in their desiccated tissues and this might explain the suggested role of ELIPs in

protection of photosystem during desiccation.


1.8 TRIPOGON LOLIIFORMIS (F.MUELL.) C.E.HUBB. A NATIVE

AUSTRALIAN RESURRECTION GRASS

1.8.1 Australian resurrection plants

Around 25 species of resurrection plants have been identified in Australia

(Gaff and McGregor, 1979). These plants can be found in all parts of Australia and in

every habitat. However these plants tend to be found mostly in shallow soils in areas

that experience intense and/or prolonged dry seasons, even if the area has high

annual rainfall (Gaff, 1981). The most suitable areas for resurrection plants in

Australia are in subtropical, most of the tropical and winter rainfall zone of South

Australia (Gaff, 1981).

Around 16 species of Australian resurrection plants are angiosperms (Gaff and

Latz, 1978). They occur in four families and eight genera (Table 1) which all but one

(Boea hygroscopica) species are monocots (Gaff, 1981). Therefore desiccation-

tolerance has probably evolved independently a number of times in Australia (Gaff,

1981).


Table 2: The occurrence of resurrection plants in various genera and families

among Australian angiosperms (Gaff, 1981).

The number of angiosperm resurrection species in various

genera and families in Australian

Family Genus

Number of

resurrection

species

Gesneriaceae

(Dicotyledonous) Boea 1

Boryaceae Borya 3

Cyperaceae

(Sedges) Fimbristylis 2

Poaceae

(True Grasses)

Eragrostiella 1

Micraira 6

Microchloa 1

Sporobolus 1

Tripogon 1

Total

4 8 16

Resurrection capability can be seen in five families and eight genera among 16

angiosperm species.


1.8.2 Tripogon Roem. & Schult a genus of true grasses

Tripogon Roem. & Schult (Poaceae: Chloridoideae) is a genus of true grasses

with approximately 40 recognised species in the world (Newmaster and Ragupathy,

2010;Peterson et al., 2010). Tripogon species are found mostly in tropical to

subtropical parts of Asia, Australasia, North America, South America and more

commonly in Africa and India (Phillips and Launert, 1971;Clayton et al., 2010

onwards). Desiccation-tolerant species of Tripogon have been found in: South

America (1), Africa (1), Malagassy, India and Australia (1) (Gaff, 1987) although it

seems no species of Tripogon genus has been identified as desiccation sensitive

(personal communication with Professor Donald Gaff). The only species of Tripogon

in Australia is T. loliiformis (F.Muell.) C.E.Hubb which is also a resurrection grass

(Gaff and Latz, 1978;Botanic Gardens Trust, 2011). Although most resurrection

plants are not agriculturally important T. loliiformis is palatable to stock (Gaff,

1981;Tothill and Hacher, 1996).

Some micro-morphological and internal structural characteristics of the genus

Tripogon have been identified in a few previous studies. Watson and Dallwitz (2004)

have recorded that on the abaxial leaf surface of Tripogon, the following structures

occur; long cells, stomatal complex, micro hairs and saddle-silica cells. They also

refer to the absence of papilla. However, these details were not conclusive based on

the micro-morphological observation that was performed in the first part of this

project. For example, notable gland cells that exist on the abaxial surface of T.

loliiformis had not been mentioned.

Micro-morphological description of the adaxial surface of some species of

Tripogon has only been mentioned in Rúgolo de Agrasar and Vega (2007). This

description is limited to the presence of epicuticular wax, long cell, macro hair,

prickle and short cell. This description lacks the existence of structures such as

stomatal complex and gland that have been observed on the adaxial surface of T.

loliiformis. This data did not cover the differences between the structures of tip to

base of the leaf. A lack of comprehensive detailed description of the micro-

morphological structure for Tripogon can be recognised in the literature.

Some internal structures of the Tripogon has been described by Watson and

Dallwitz (2004) for example; Tripogon has Kranz type XyMS+ (double sheath)

bundle structure (Figure 2) (Lang et al., 2004). XyMS+ (“double sheath") species


have a layer of mestome sheath (inner bundle) under the bundle sheath.

Physiologically it is known that this genus is a C4 grass with NAD-ME biochemical

structure (Figure 3) biochemical type (Anderson and Briske, 1990;Lang et al., 2004).

In NAD-ME species, carbon is fixed in aspartate in the mesophyll then aspartate is

decarboxylated by NAD-malic enzyme in the bundle sheath cell mitochondria, and

alanine is returned to the mesophyll cells for the next cycle. Tripogon has been

described to be diploid with 20 chromosomes (2n=20; x=10) (Kitajima and Butler,

1975;Lang et al., 2004).


Figure 3: A schematic demonstration of enzyme compartmentalisation and

structural characteristics in C4 biochemical NAD-ME type photosynthesis.

In NAD-ME species, Carbon is fixed in aspartate in the mesophyll then aspartate is

decarboxylated by NAD-malic enzyme in the bundle sheath cell mitochondria, and

alanine is returned to the mesophyll cells for the next cycle. CO2 released in bundle

sheath is used in Calvin cycle for producing carbohydrate mediated by Rubisco

enzyme inside chloroplast. Modified from (Jami et al., 2007).

Figure 2: A schematic structure of C4 (Kranz)

type XyMS+.

In C4 XyMS+ species the procambium gives rise

to xylem, phloem, and a mestome sheath;

associated ground meristem differentiates into C4

mesophyll tissue and the bundle sheath. XyMS+

(“double sheath") species have a layer of

mestome sheath (inner bundle) under bundle

sheath. Bundle sheath (bs); Mesophyll (m);

Mestome sheath (ms); Metaxylem vessel (mv);

Phloem (p); Xylem (x).


1.8.3 Tripogon loliiformis (F.Muell.) C.E.Hubb an Australian resurrection grass

Tripogon loliiformis (Five-minute Grass, Rye Beetle-grass) is a small tufted

annual to short-lived perennial grass native to Australia and New Guinea (The Plant

List, 2010;Clayton et al., 2010 onwards). In Australia it grows almost everywhere

except Tasmania, from tropical forests, temperate climates to arid and semi-arid

areas of central Australia (Figure 4) (Orchard and Wilson, 2005;Botanic Gardens

Trust, 2011). It grows in a wide range of habitats such as: rocky slopes, plateaux and

outcrops of granite and sandstone, on plains in red sand or sandy to clayey loams in

open Acacia woodlands especially mulga and flowers and fruits throughout the year

(Orchard and Wilson, 2005).

Figure 4: Distribution of Tripogon loliiformis in Australia (Orchard and Wilson,

2005).

Dots illustrate geographical locations of Tripogon loliiformis herbarium samples

collected in Australia.

Tripogon loliiformis has small leaves, 1-7.5cm long and 0.5-1.5 mm wide

which usually are bearded with a cilicate membrane ligule (0.1-2 mm); the flowering

culms range from 3.5-20 (-55) cm high with inflorescence composed of racemes

(Orchard and Wilson, 2005;Clayton et al., 2010 onwards). This plant is very

polymorphic particularly in the hairiness of the leaves and the size of the plant (Gaff

and Latz, 1978;Gaff, 1981). However all tested specimens have shown resurrection

capability (Gaff, 1981).


Resurrection characteristics of T. loliiformis and other Australian

resurrection plants

Tripogon loliiformis is considered a resurrection grass as its vegetative tissues

can withstand air-dry state (Figure 5). Our knowledge about T. loliiformis and other

Australian resurrection plants is mainly limited to species identification and

distribution. There have also been some studies on general anatomical and

physiological resurrection characteristics of these plants such as pigmentation,

photosynthesis status and changes in nitrogen content (Gaff and Latz, 1978;Gaff and

McGregor, 1979;Gaff, 1981). Among Australian resurrection plants, genus Borya

has received more attention (Gaff and McGregor, 1979;Gaff, 1981). There is a lack

of information about the roots structural responses and some anatomical responses

such as the change in epidermis, stomata, xylem and vacuole during dehydration.

The known dehydration responses of T. loliiformis are limited to its degree of

dehydration survival (Gaff and Latz, 1978) and its quick rehydration response upon

supply of water (Tothill and Hacher, 1996). Some of its physiological responses

during dehydration and rehydration are also known, such as: chlorophyll retention

(homoiochlorophyllous) (Gaff, 1981), production of pigment (Gaff, 1981) and

change in nitrogen level (Gaff and McGregor, 1979) during dehydration.


Figure 5: The top panel shows Tripogon loliiformis in its natural habitat while

the bottom panel shows this plant grown from seeds in glasshouse in in 65mm

pots.

(A) and (D) hydrated, (B) and (E) dehydrated and (C) and (F) rehydrated plants.


Tripogon loliiformis as an experimental model

Tripogon loliiformis has many characteristics which make it suitable to be used

as an experimental model plant for studying dehydration-tolerance. Some of these

characteristics are: 1) it exhibits most of the typical elements of resurrection plants,

including the ability to withstand severe desiccation in vegetative tissues, pigment

production during desiccation and quick rehydration (Gaff and Latz, 1978;Gaff,

1981), 2) it grows almost everywhere on the Australian mainland and grows in a

wide range of habitats (Orchard and Wilson, 2005), 3) it can reach maturity and

produce seeds in a relatively short period of time, 4) it can be grown relatively easily

under greenhouse conditions (Gaff, personal communication with Mundree), 5) it is

in the same family (Poaceae) as true cereals which are an important source of staple

food and 6) it is a diploid plant which makes it easy for molecular work.

Investigation of anatomical, morphological and molecular behaviour in T.

loliiformis as a model plant during different stages of dehydration and rehydration

would result in a better understanding of dehydration-tolerance mechanisms and the

interaction of each part with one another (from gene expression to physiological

responses and anatomical changes). This knowledge can provide a better insights

into the mechanism of dehydration-tolerance which could eventually contribute to

the development of drought-tolerant crops.

1.9 BAG, A FAMILY OF PRO-SURVIVAL PROTEINS

Dehydration in plants normally results in activation of programmed cell death

(PCD), however the vegetative tissues of resurrection plants avoid PCD, suggesting

that they might be utilising pro-survival (anti-apoptotic) molecules in order to

prevent apoptosis. The bcl-2-associated athanogene (BAG) family are considered

molecular chaperones and have anti-apoptotic (or pro-survival) properties, however,

their function is not limited to this. BAG proteins are an evolutionary conserved

family of multifunctional proteins. BAG proteins have been found in many

organisms from yeast to plants and animals (Doukhanina et al., 2006). These proteins

share a conserved region near the C terminus, known as the BAG domain containing

110–130 amino acids. The BAG domain interacts with the ATPase domain of heat-


shock protein Hsp70 (Hsc70) however, the N-terminal region is different among

different BAG members (Brive et al., 2001;Sondermann et al., 2001).

1.9.1 In animals

The first mammalian BAG (BAG1) was identified in a cDNA library screening

of mouse embryo designed to identify the binding partner of Bcl-2 using human Bcl-

2 as bait (Takayama et al., 1995). BAG1 was shown to have anti-apoptotic properties

suggesting it is involved in PCD. Later studies revealed the BAG family are

molecular chaperones (Briknarová et al., 2001). In humans, six members of the BAG

family (BAG1-6) have been identified, acting as chaperone regulators interacting

with Hsp70 chaperone proteins and form complexes with a variety of transcription

factors leading to regulation of a number of diverse processes ranging from

apoptosis, tumorigenesis, neuronal differentiation, stress responses, and the cell cycle

(Zeiner and Gehring, 1995;Wang et al., 1996).

1.9.2 In plants

Although BAG proteins had been extensively studied in animals, plant BAG

family had been unknown until 2003. This is at least partly due to the problems with

identification of the BAG family in plants. Plant cells can undergo programmed cell

death (PCD), the primary sequence of plant BAG does not have strong similarity with

animal BAG homologues and therefore it was not possible to be found using usual

bioinformatics tools. It has been suggested that this difference is mainly due to the

difference between animal and plant PCD regulators (Doukhanina et al., 2006). The

first identification of plant BAG family was by Yan et al. (2003) suggesting eight

proteins for Arabidopsis BAG family. Later this classification was refined by

Doukhanina et al. (2006) using bioinformatics functional similarity approach

[profile-sequence (PFAM) and profile-profile (FFAS) algorithms] that identified

seven homologs of BAG family in Arabidopsis (AtBAG1-7). Four of these have

domain organisation similar to animal BAGs (Figure 5) (Doukhanina et al., 2006).

Transgenic tobacco plants overexpressing AtBAG4 showed resistance toward a

number of abiotic stresses (Doukhanina et al., 2006). In another study by Hoang

(2014) rice plants overexpressing AtBAG4 and OsBAG4 (from rice) were shown to

be more tolerant to salinity stress than the wild type Nipponbare rice variety.


AtBAG6 was first identified in a calmodulin binding protein screening (Kang et al.,

2005) and plays a role in plant defence to necrotrophic pathogens. Williams et al.

(2010) revealed that the BAG family in Arabidopsis is very diverse and plant BAGs

have a cytoprotective role. That study also demonstrated that AtBAG7 is localised in

the endoplasmic reticulum (ER) and interacts directly in vivo with AtBiP2 (an ER

localised HSP70) which is a molecular chaperone. The same study demonstrated

AtBAG7 is an essential component for the proper maintenance of the unfolded

protein response (UPR) during heat and cold-tolerance. While plant BAGs, and

AtBAG4 in particular, have shown to be promising for conferring abiotic stress in

transgenic plants, plant BAGs are still not well characterised.

1.10 CONCLUSION, AIMS AND OBJECTIVES

Although molecular, physiological and to a lesser extent, structural responses

of resurrection plants to dehydration have been studied since 1912 (Irmscher), most

research has been limited to a small number of species, and these have been mainly

from southern Africa (Table 1). Studying the resurrection behaviour of other species

can provide us with new insights to understand this unusual behaviour. Tripogon

loliiformis (F.Muell.) C.E.Hubb is indigenous to Australia and possesses

characteristics to be considered as a good model plant for studying resurrection

behaviour. This PhD project has examined this species as a model for studying the

resurrection behaviours during dehydration and rehydration.

1.10.1 Aims

Based on the knowledge gap in the field the main aim of this project is to

address the following question:

What are the desiccation-tolerance mechanisms of Tripogon loliiformis

(F.Muell.) C.E.Hubb?

Some sub-questions that are raised by the main question are as follows:

1. What is the general hydrated leaf structure of T. loliiformis?


2. What are the structural changes of T. loliiformis during various stages of

dehydration and rehydration?

3. What are the physiological changes of T. loliiformis during various stages of

dehydration and rehydration?

4. What is the functional mechanism of BAG4?

5. What is the difference between the function between BAG4 from a

resurrection plant with a desiccation-sensitive plant?

1.10.2 Objectives

The objectives for this project based on the project questions identified above can

be formulated as:

1. Investigation of general micro-morphological and anatomical structure of T.

loliiformis hydrated leaves.

2. Investigation of micro-morphological and anatomical changes of T. loliiformis

during various stages of dehydration and rehydration.

3. Investigation of physiological responses of T. loliiformis during various stages

of dehydration and rehydration.

4. Investigation into protein-protein interaction of BAG4.

5. Investigation into the difference between the protein-protein interaction

between BAG4 protein from a resurrection plant and a desiccation-sensitive

plant.


Chapter 2: General Materials and Research Design 41

Chapter 2: General Materials and Research

Design

2.1 RESEARCH DESIGN

The objectives of this research project (discussed in section 1.10) can be

categorised under three main areas; structural, physiological and molecular

approaches. Together these approaches provide a detailed picture of resurrection

behaviour in Tripogon loliiformis during dehydration and rehydration. Likewise the

methods used in this experiment can be divided into three general categories.

2.1.1 Structural experiments

A combination of histological techniques were used to study the structure and

structural changes in T. loliiformis leaves. As the structure of this plant is not well

described particularly on adaxial surface, first surfaces as well as the internal

structures of this plant were observed from tip to the base of the leaves. Then

desiccated leaves were observed in order to study the structural changes during

desiccation.

One of the main challenges of studying structural changes during dehydration

and rehydration is to observe the plant tissues in their natural state. Conventional

methods of tissue preparation for internal plant structure investigation usually

involve possible tissue altering procedures such as fixation, hydration, dehydration,

infiltration, embedding in a solid matrix, sectioning and staining (Bewley, 1979).

Any changes in hydration level of samples result in artificial structure changes of

tissue. In order to observe the structure of the tissues in their closest to natural state,

techniques were optimised to suit the small and delicate leaves of T. loliiformis and

to have minimum impact on the hydration level of the tissue. These techniques

included techniques such as an improved freehand sectioning method developed for

sectioning of small, delicate plant materials, and environmental scanning electron

microscopy (ESEM) where plant tissue can be placed in the chamber (which filled

with air) without being fixed nor coated and can be observed in its natural hydration

state. The following flowchart demonstrates an overview of experimental design for

42 Chapter 2: General Materials and Research Design

the structural studies of this research project. The techniques used in the structural

studies are described in detail in section 3.2.


Structural studies

General features Structural changes

External (morphology)

Internal (anatomy)

Whole leaf (folding)

Microscopic (cellular level)

Light microscopy Epidermal replica Freehand sectioning Environmental scanning electron microscopy (ESEM)


2.1.2 Physiological experiments

The physiological observations were design to determine the integrity of the

cell during dehydration and rehydration, as well as observing the main physiological

changes during dehydration and rehydration. For determination of the cell integrity,

two main areas were monitored; 1) chlorophyll integrity and 2) the level of

membrane integrity. For measuring the level of chlorophyll integrity two parameters

were quantified; 1) chlorophyll fluorescence and 2) chlorophyll content. Chlorophyll

fluorescence was measured using the two parameters of maximum photochemical

efficiency (Fv/Fm) and quantum yield of PSII (ΦPSII) (discussed in section 4.3.2)

using MINI-PAM. Membrane integrity was assessed using two methods of

electrolyte leakage using a conductivity meter and propidium iodide staining.

The main physiological changes among resurrection plants that have been

shown to be involved are pigmentation and gas exchange. Physiological observations

of this project examined the changes in photosynthesis rate and pigmentation using

hydration level as reference. For measuring the photosynthetic rate CO2 assimilation

of the whole plant was measured during dehydration and rehydration (using LI-

COR). The accumulation of the pigmentation was observed using light microscopy.

The following flowchart demonstrates an overview of experimental design for

physiological studies part of this research project. The techniques used in the

molecular studies are described in detail at section 4.2.


Physiological studies

Cell integrity Responses

Chlorophyll integrity

Membrane integrity

Photosynthetic rate

Hydration level

Chlorophyll fluorescence

Chlorophyll content

Propidium iodide Staining

Electrolyte leakage

Relative water content

CO2 assimilation rate

Fresh dry weight ratio


2.1.3 Molecular experiments

The possible role of anti-apoptotic proteins in desiccation tolerance has not

been studied in previous research on desiccation-tolerant plants. In order to address

this, we focussed on understanding the role of bcl-2-associated athanogene 4

(BAG4), an anti-apoptotic protein, and its possible involvement in desiccation

tolerance. For addressing the objectives 4 and 5 (section 1.10.2), the molecular

studies part of this research project looked at one of these proteins called bcl-2-

associated athanogene 4 (BAG4). The molecular mechanism of this protein function

is still unknown. In order to understand the possible involvement of this protein in

desiccation-tolerance, first the function of this protein needed to be investigated. In

order to understand the function of BAG4, the proteins which interact with this

protein were identified to identify the pathway it is involved. After understanding the

pathway that this protein is involved in, the proteins that interactive with BAG4 from

T. loliiformis (as a desiccation-tolerant plant) were compared with proteins that

interacted with BAG4 from Arabidopsis thaliana (as a desiccation-sensitive plant).

This was done in order to understand the possible difference between the mechanism

this anti-apoptotic protein works in a resurrection plant as compared with a

desiccation-sensitive plant. The protein-protein interaction was investigated using a

protein microarray.

The technical procedures of the molecular studies can be divided into five main

areas; 1) protein expression, 2) protein purification, 3) verification, 4) protein

microarray and 5) data analysis. Proteins expression involved insertion of the BAG4

protein from A. thaliana (AtBAG4) and T. loliiformis (TlBAG4) in the expression

vector which was facilitated using TOPO® Cloning technique and transient

expression of the constructs in the leaves of Nicotiana benthamiana using

Agrobacterium-infiltration method. Protein purification involved extraction of the

total protein from infiltrated leaves and purification of the protein using IgG

sepharose beads. The verification techniques included PCR, agarose gel

electrophoresis, Mini prepping, transforming Escherichia coli and Agrobacterium

tumefaciens using heat shock and electroporation transformations, restriction enzyme

digestion, sequencing, histochemical GUS assay were used to determine the protein

expression, and the success of protein purification was assessed using SDS PAGE,

Coomassie blue staining and Western blotting techniques. Protein microarray


involved hybridisation of the extracted proteins to the microarray chips and scanning

the hybridised chips using a GenePix scanner. Finally the statistical and

bioinformatics analysis were used to analyse the data. The techniques used in the

molecular studies are described in detail in section 5.2.


Molecular studies

Protein expression

Data analysis

Vector constructio

n

Protein purification

Verification of results

Protein microarray

Transient expression

Protein extraction

IgG bead purification

Protein expression

Protein purificatio

n

Microchip Hybridisatio

n

Microchip scanning

Statistical analysis

Bioinformatics analysis

TOPO® cloning in

pENTR

LR Cloning in pYL436

Agro-transformation

Agro- infiltration

PCR Total protein extraction

IgG bead separation

Cleavage purification

Sequencing

Mini prepping

Restriction digestion

Histochemical

GUS assay

SDS PAGE

Coomassie blue staining

Western blotting

Microchip Hybridisation

Microchip scanning

Generating report file

Determining significant

signals

GO enrichment

Determination of significant protein

interactors

FDR


2.2 GENERAL MATERIALS

2.2.1 Sources of specialised reagents

All the reagents used in this project were sourced from reputable scientific

supply companies such as Sigma Aldrich (Australia), Roche Diagnostics

(Switzerland) and Crown Scientific (Australia). IgG Sepharose beads were purchased

from GE healthcare Life Sciences (Australia) (product number 3420), Precession

protease was purchased from Amersham Biosciences (Sweden) (product number 27-

0843-01), c-Myc Monoclonal Antibody from Invitrogen (product number AHO0062)

and CY5-conjugated anti-mouse IgG from Invitrogen (Australia) (product number

R950-25).

2.2.2 Oligodeoxyribonucleotide synthesis

Oligonucleotides were synthesised by GeneWorks (QUT, CTCB) in a

concentration of 100 µM. The stock primers were diluted to 20 µM before PCR.

2.2.3 Bacterial strains

Agrobacterium tumefaciens strain AGL1 was used for inoculation of the plants

and Escherichia coli strain XL1-Blue was used for general plasmid cloning.

2.2.4 List of general solutions

EDTA: Ethylenediaminetetraacetic acid, pH 8.0

Luria-Bertani (LB) liquid growth media: 1 % (w/v) bacto-tryptone, 0.5 %

(w/v) bacto-yeast extract, 170 mM sodium chloride.

Luria-Bertani (LB) agar: LB liquid growth media solidified with 1.5 %

bacto-agar.

TE Buffer: 10 mM Tris-HCl, pH 8.), 1 mM EDTA.

Agarose gel loading dye (6X): 0.25 % (w/v) bromophenol blue, 50 % TE, 50

% glycerol

Propidium iodide (PI): 2.5 µg/mL in water

Tris-buffered saline (TBS) buffer: 50 mM Tris, 150 mM NaCl


Washing buffer: 50 mM Tris HCl pH 7.5, 150 mM NaCl, 10 % Glycerol, 0.1

% Triton

Infiltration media: 10 mM MgSO4.7H2O, 9 mM MES, pH 5.6

Cleavage buffer: 50 mM Tris HCl pH 7.0, 150 mM NaCl, 1 mM EDTA, 1

mM DTT, 0.1 % Triton

SDS running buffer: 250 mM Tris, 1.92 M glycine, 1 % SDS

SDS sample loading buffer: 50 mM Tris (pH 6.8), 1 % SDS, 15 % Glycerol,

0.025 % Bromophenol blue, 10 mM DTT.

Super Optimal Broth (SOB/SOC medium): 2% vegetable peptone, 0.5%

yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2, 10 mM MgSO4, 20 mM

glucose

TAE buffer: 0.04 M Tris – Acetate, 0.0001 M EDTA, pH 8.0

TTBS buffer: 50 mM Tris, 150 mM NaCl, 0.05 % Tween 20, pH 7.6 (adjusted

by HCl)

Blotting buffer: 25 mM Tris, 192 mM glycine, 10 % methanol

Blocking buffer: TTBS, 5 % (v/w) skim milk powder

Ponceau S staining solution: dH2O, 1 % (v/v) acetic acid (16.6 M), 0.5 %

(w/v)

2.2.5 Plant material

Tripogon loliiformis plants were collected from Charleville (GPS: S 26.42686.

E 146.25002) an arid area in the state of Queensland, Australia. Inbred seeds derived

from a single mother plant were collected and stored in the fridge until being used for

all the experiments (Queensland Herbarium voucher accession number: Williams

01). Seeds were germinated in a mixture of red soil and seedling germination media

(50 % by volume each) in 50 mm pots in relatively close density and were grown in

growth cabinets under a 12h photoperiod (cold white fluorescent light, light intensity

of 900±100 µmolm-2

s-1

) with day-night temperatures of 30-20C and watered two to

three times a week with tap water and fertilized once a month. Dehydration and


desiccation was induced through withholding the water until plants reached an air-

dry state before rehydration through rewatering.


Chapter 3: Tripogon loliiformis Displays Structural Features and Changes that Protect It During Dehydration 53

Chapter 3: Tripogon loliiformis Displays

Structural Features and

Changes that Protect It During

Dehydration

3.1 INTRODUCTION

Leaf structure has been shown to play a significant role in plant stress

tolerance, especially drying (Gutschick, 1999). In this research project leaf structural

aspects are divided into two categories, 1) structural features and 2) structural

changes. Structural features such as leaf shape, trichomes and cuticle could be

permanent or as a response to environmental conditions, growing gradually and are

irreversible. Structural changes such as leaf folding are rapid responses to

environmental changes, quick and quite often are reversible. Plants structural

features and changes are to minimise radiation damage or reduce water loss rate

(dehydration avoidance).

3.1.1 Structural features

The tissues of desiccation-tolerant and -sensitive plants share many structural

aspects; however, there are structural changes unique to viable leaves of desiccation-

tolerant plants. These unique structural changes occur mainly at the cellular level and

are not observed in the leaves of desiccation-sensitive plants. These changes take

place during very low hydration levels and are believed to be mainly for maintenance

of cell membrane integrity and prevention of mechanical damage during extreme

dehydration (Farrant, 2000;Vander Willigen et al., 2003;Vander Willigen et al.,

2004). Some of these structural changes have, also, been observed in the seeds of

orthodox seeds.

3.1.2 Structural changes

Leaf folding and rolling remain the most obvious structural changes during

dehydration which have been reported in a number of resurrection plants (e.g. Gaff,

1981;Sherwin et al., 1998;Vander Willigen et al., 2003;Moore et al., 2007;Georgieva

et al., 2010). Leaf folding and rolling result in reduced leaf surface area. Reduced

54 Chapter 3: Tripogon loliiformis Displays Structural Features and Changes that Protect It During Dehydration

leaf surface area is a mechanism used by plants that reduces dehydration rate, as well

as light-induced damage during dehydration and desiccation.

One of the major problems that resurrection plants face during extreme

dehydration is the extensive reduction in cell volume. Extensive shrinkage in cell

volume can lead to fusion of the cell membrane as the result of membrane folding on

itself or plasma membrane rupture as the result of tension between plasma membrane

and plasmodesmatal connections against the rigid cell wall (Cosgrove,

2000;Hoekstra et al., 2001). To minimize these mechanical stresses, resurrection

plants respond by structural changes at the cellular level. Cell wall folding and

vacuole fragmentation are two of the main such changes observed in several

resurrection plants (Iljin, 1957;Vertucci and Farrant, 1995;Farrant, 2000;Vander

Willigen et al., 2004).

In order to investigate the structural aspects of Tripogon loliiformis, the

research was designed to first build an understanding of general morphological and

internal structures (anatomy) of the leaf from apex to base in the hydrated state; after

which structural changes of the leaves in the desiccated state were investigated.

The two main objectives of research performed in this chapter were;

1. To characterise the general structural features of T. loliiformis leaves at the

hydrated state.

2. To determine the leaf structural changes during dehydration and desiccated.

3.2 MATERIALS AND METHODS

3.2.1 Plant materials

The third fully developed leaf of three month old plants was used in all

experimental work. Seven biological replicates were collected for each structural

experiment (n = 7). Separate sets of plants were used for each experiment due to the

destructive nature of the sampling process. Internal and external structures of apex

(top 5 mm), middle and base (bottom 3-5 mm) of leaves were observed at hydrated

and desiccated state for structural studies.


3.2.2 Methods

Internal structures, external structures and structural changes of T. loliiformis

leaves were investigated using a combination of well-established histological

methods, namely epidermal replica construction, freehand sectioning, light

microscopy and environmental scanning electron microscopy (ESEM) and

computerised data processing analysis.

Freehand sectioning

In order to observe the internal structures of the desiccated leaves in their

natural state, a modified freehand sectioning method was developed. This optimised

method requires the sample to be placed horizontally on a flat surface. Razor blade

was held in the right hand (if right handed) and the left hand was used to stabilise the

sample. Left hand was rested on the surface close to the sample with the index finger

holding the sample. The razor blade rested perpendicular to the fingernail and the

fingernail of the left hand alone determined and controlled which site on the sample

were cut and how thin the slice were taken. Sections were directly mounted onto a

glass slide in a drop of water. Samples were covered with a coverslip without

staining or processing. Sections from dry samples were mounted directly in

anhydrous DPX mounting agent to avoid rehydration. These sections were examined

under a Nikon Eclipse 50i light microscope with an attached Nikon DS-Fi1 camera

head and photographed at different magnifications. Two sections from the apex,

middle and the base of leaf blades for each biological replicate were used for data

analysis.

Epidermal replica production

Epidermal replica production (Hilu and Randall, 1984) was performed on

leaves of T. loliiformis by coating the surfaces of the leaf with a thin layer of clear

nail polish. After nail polish was dry, the dry nail polish surface (nitrocellulose) was

peeled using sticky tape. The tape was then placed onto a glass slide. The epidermal

layer was made visible by light microscopy.

Environmental scanning electron microscopy (ESEM)

Unprocessed freshly harvested leaves were used for ESEM. To minimise

dehydration a cold stage was used to bring the sample temperature down to 2C.


Double-sided sticky tape was used to mount the samples on the cold stage. For

observations of leaf folding and rolling during dehydration, ESEM was used, fresh

leaves were cut and mounted vertically on Blu-Tack® mound. This was placed over a

glass slide which was fixed to a cold stage (2˚C) in ESEM chamber using double-

sided sticky tape. The upper part and base of seven leaves were used for observation

of leaf folding and rolling respectively. All the ESEM observations were surveyed

using a FEITM

Quanta 200 ESEM.

Quantitative analysis of microscopy data

The density of specialised structures such as stomatal complexes, prickle hairs

and glands were calculated from the middle region of both abaxial and adaxial leaf

surfaces (Chen et al., 2001). Seven biological and two technical replicates were

randomly selected and photographed for observation of each region. Computer-

aided determination of structure counts and distance measurements were performed

using Nikon NIS Elements Basic Research software (Version 3.21).

3.3 RESULTS

3.3.1 General structural observations

Leaf structural features play a critical role in stress tolerance. General structural

observation was done using techniques including ESEM, freehand sectioning, light

microscopy, epidermal replica construction and computerised data processing

analysis using fresh T. loliiformis leaves. A number of cells and structures of the leaf

were identified from observation of internal and external structure of T. loliiformis

(Table 3).

Investigation into leaf anatomy of T. loliiformis

The observation of internal structures of the T. loliiformis leaves from apex to

the base revealed that the number of vascular bundles was found to increase from the

apex to the base of the leaf (3-11, respectively). On the adaxial surface, the

intercostal regions (the region between bundles) were composed of enlarged, thin-

walled bulliform cells (Figure 6A,B). These bulliform cells formed parallel stripes

between the vascular bundles (in intercostal regions), which in transverse section


forms groups of fan-shaped cells which in turn forms grooves during dehydration

(Figure 6C,D).


Table 3: Abbreviations and location/section of Tripogon loliiformis leaf

structures

Cell and Structure Type Abbreviation Location/section

Bulliform Cell bu Adaxial, Transverse

Bundle Sheath bs Transverse

Colourless Cell co Transverse

Epicuticular Wax ew Adaxial

Gland g Adaxial, Abaxial

Guard Cells gu Adaxial, Abaxial, Transverse

Hooked Trichome ho Adaxial, Abaxial, Transverse

Long Cell lo Adaxial, Abaxial, Transverse

Mesophyll m Adaxial, Abaxial, Transverse

Mestome Sheath/Inner Bundle Sheath ms Transverse

Metaxylem Vessel mv Transverse

Papilla pa Adaxial, Abaxial

Phloem p Transverse

Prickle-Hair pr Adaxial

Prickle-Hair Basal Cell pr-bc Adaxial

Sclerenchyma s Transverse

Short Cell sh Adaxial, Abaxial

Short Cell- nodular sh-n Adaxial, Abaxial

Silica Cell- saddle si-s Adaxial, Abaxial

Stomatal Complex st Adaxial, Abaxial, Transverse

Subsidiary Cell su Adaxial, Abaxial, Transverse

Xylem x Transverse


Vascular bundles demonstrated typical Kranz anatomy associated with C4

plants and characterised by XyMS+ (mestome present) (Figure 6B). The

chloroplasts in the bundle sheath cells exhibited centripetal distribution (Hatch et al.,

1975) (Figure 6A). Vascular tissue is surrounded by mestome sheath (also known as

inner bundle sheath), bundle sheath and mesophyll (Figures 6B and 10A). Bundle

sheath is a single layer of thick wall, large cells; however in the base of the leaf,

bundle sheath can have more than one layer of cells on adaxial side (data not shown).

On both the abaxial and adaxial sides, bundle sheath and mesophyll cells were

interrupted by sclerenchymatous tissue, except on the adaxial side of the apex and

middle regions of the leaf (Figures 6B, 9abc and 10A,B). A thick cuticle layer was

observed on both surfaces except over bulliform cells where it appeared as a thin

layer (Figure 10A-C).

Investigation into micro-morphological leaf structures of T. loliiformis

Micro-morphological structures of T. loliiformis leaves were observed using

epidermal replica production and ESEM. Previous studies have poorly described the

abaxial surface of Tripogon (Oliver et al., 2000a;Lang et al., 2004) with adaxial

surfaces of Tripogon have not been described in the literature. Adaxial and abaxial

surface structure of T. loliiformis leaves are remarkably different (Figure 7). There

are a number of features on the adaxial surface which do not exist on the abaxial

surface, namely bulliform cells, prickle-hairs, papillae and epicuticular wax.

Furthermore the structure from apex to base changes. For example there is a row of

hooked trichomes on midvein of the abaxial side of the apex which does not exist on

other parts of the leaf surfaces (except the edges of the leaf).


Figure 6: Freehand transverse sections of hydrated and dehydrated leaves of

Tripogon loliiformis leaves.

(A) and (B) hydrated and (C) and (D) dehydrated leaves. Arrows are pointing at the

pigmented abaxial surface of desiccated leaves. Bulliform cell (bu); Bundle sheath

(bs); Mesophyll (m); Sclerenchyma (s); Phloem (p) and Xylem (x). Scale bar at (A),

(B), (C) and (D) = 100 µm.


Figure 7: Environmental scanning electron microscopy (ESEM) images of

representative hydrated leaves of Tripogon loliiformis.

(A) and (B) show the adaxial surface and (C) and (D) show the abaxial surface.

White lines on the top of (A) and (C) indicate the position of side veins and black

lines indicates the position of midveins. Adaxial surface is covered with a thick layer

of epicuticular wax (B). Bulliform cell (bu); Gland (g); Long cell (lo); Papilla (pa);

Prickle hair (pr); Prickle-hair basal cell (pr-bc); Saddle silica-cell (si-s); Stomatal

complex (st); Nodular short-cell (sh-n). Scale bar (A) and (C) = 100 µm and in (B)

and (D) = 25 µm.


Adaxial surface

The adaxial surface structure appeared to be more complex than the abaxial

surface; and additionally the surface structure of the midvein differed significantly

from apex to the base of the leaf (Figures 7 and 9). Stomatal complexes were

organised in longitudinal rows parallel to the veins between the costal and intercostal

regions (Figure 7). However, on the adaxial surface of the midvein the stomatal

complexes occurred only on the costal regions of the upper part of the leaf (Figure

7A). The density of stomatal complexes was approximately 25 % higher on the

adaxial surface in comparison to the abaxial surface (Table 4).

Table 4: The density of stomatal complex, gland and prickle hair on adaxial and

abaxial surface of the leaf per mm2.

Stomatal

complex

Gland Prickle

hair

Adaxial 403±82.2 49.7±21.4 80.1±25.8

Abaxial 322.2±66.4 75±20.4 Absent

These numbers are means of 7 replicates with two reading from each replicate.

Three types of trichomes were observed on adaxial surface of T. loliiformis

leaves; namely prickle hair, macro-hair and gland. Prickle hairs occurred only on the

costal regions of the adaxial surface as a replacement to the saddle silica-cells

(Figure 7A,B). Prickle hairs are absent on the costal region of the midvein in the

apical and mid regions of leaves, where saddle silica-cells are absent (Figure 7A).

Macro-hairs are usually abundant on the adaxial surface, however, plants that were

grown under high humidity did not develop macro-hairs. The glands were of typical

Chloridoideae two-celled salt glands (Liphschitz and Waisel, 1974;Amarasinghe and

Watson, 1988) (Figure 7A). The gland density on the abaxial surface was

approximately 50 % higher than the adaxial surface (Table 4).

In T. loliiformis, short cells consisted of saddle silica-cells alternating with

nodular short-cells; no cork cells were however observed (Figure 7). On the adaxial

surface, each costal region was covered with one row of short cells (Figure 7A,B)

except on the midvein in the apical and mid regions of the leaves (Figure 7A). Long


cells (also called fundamental cells) dominated the costal regions of the adaxial

surface, and were covered with papillae (Figure 7A,B).The adaxial surface of the

leaves was covered by a thick platelet-type epicuticular wax based on the

classification of epicuticular wax by Barthlott et al. (1998) (Figure 7B). The non-

orientated platelets epicuticular wax crystalloids were well developed and covered all

cells.

Abaxial surface

Similar to the adaxial surface, stomatal complexes on abaxial surface of the T.

loliiformis leaves were organised in longitudinal rows parallel to the veins between

the costal and intercostal regions (Figure 7C,D). Salt glands were observed in the

intercostal regions of the abaxial surface (Figure 7C). Macro-hairs on the abaxial

surface were only observed in plants that were grown under low humidity (data not

shown). On the abaxial surface, there were two or more rows of short cells on each

vein (Figure 7C,D). Long cells made up the greatest proportion of the epidermal

cells were present on the abaxial surface. No papillae and prickle hairs were

observed on the abaxial surface. Hooked trichomes were observed on the edges of

the leaf and on the midvein close to the tip of the leaf on the abaxial surface.

3.3.2 Structural changes

Previous studies have shown substantial structural changes occur in

resurrection plants during desiccation. These changes have a significant role in

desiccation-tolerance by helping reduce mechanical and oxidative stress. Some of the

reported structural changes in resurrection plants during desiccation are leaf folding

and rolling, vacuole fragmentation, cell wall folding and changes in the structure of

chloroplast and mitochondria (Sherwin and Farrant, 1998;Farrant, 2000;Vander

Willigen et al., 2003;Vander Willigen et al., 2004).

Drying and desiccated leaves of T. loliiformis were observed for structural

changes using ESEM and freehand sectioning. These changes can be classified in

two main groups of 1) leaf folding and 2) internal structural changes. Leaf folding

can be divided into two types of tight folding and loose folding (or rolling)

respectively. Cell wall folding and vacuole fragmentation have been observed as

internal structural changes.


Leaf folding

Tripogon loliiformis undergoes a range of changes during dehydration and

rehydration (Figures 8 and 10). Leaf folding is the most apparent change and occurs

at around 65 % RWC (Figures 6 and 8). At the apex and middle of the leaf, leaf

folding is tight (Figure 8A-C), In contrast, folding was loose at the base of the leaf,

folding was loose (Figure 8D-F). A comparison between the anatomy of the midvein

from the apex to the base of the leaves revealed that tight and loose folding is

directly related to the absence or presence of sclerenchymatous tissue on adaxial side

of the midvein (Figure 10a-d). When the sclerenchyma is absent at the apex and

middle of the adaxial side of the leaf, leaf folding is tight (Figures 8A-C and 9 A,B),

while in the base of the leaf where the sclerenchyma on the adaxial side is developed,

tight folding does not occur (Figure 8D-F and 9c). The existence of

sclerenchymatous tissue is directly related to the occurrence of other structures

including silica cells and prickle hairs. It was evident that whenever

sclerenchymatous tissue was present, silica cells and prickle hairs were also evident

(Figure 9).

Changes in internal structure (anatomy)

Light microscopy of untreated leaves using freehand sectioning provided insights

into the internal structure of the leaves in their natural state during dehydration and

rehydration. Almost all the internal soft structures such as bulliform cells, mesophyll,

mestome sheath and young sclerenchymatous tissue showed varying degrees of cell

wall folding during desiccation state (Figure 10B). The cell wall folding also

resulted in shrinkage of the total volume (Figure 10B). Multiple smaller vacuoles

were observed in the bundle sheath cells, which were filled with pale orange-brown

components (Figure 10B). Following rehydration, cells returned to their original

state without any apparent injury (Figure 10C).


Figure 8: Difference between the process of leaf folding in middle and base of

fully grown Tripogon loliiformis leaves using environmental scanning electron

microscopy (ESEM).

(A), (B) and (C) images of tight leaf folding process in the middle region of a leaf

and (D), (E) and (F) loose folding (or rolling) in the base of a fully developed leaf.

Scale bar = 200 µm.


Figure 9: Differences in internal and external structures from apical to basal

regions of Tripogon loliiformis leaves.

Hydrated leaves were harvested, structural features of the leaves from apex to the

base made visible using environmental electron microscopy (ESEM) (left side) and

freehand sectioning (right side). Adaxial surface apical (A), middle (B) and basal (C)

regions of the leaf. Internal structure of apical (a), middle (b) and basal (c) regions of

the leaves using freehand sectioning. The white lines represent the side veins and

black lines represent the midveins. Midvein in adaxial (A) and (B) but protruded in

(C). Arrow is pointing at sclerenchymatous tissue on adaxial surface in (a), (b) and

(c). No sclerenchymatous tissue exist on adaxial part of the midvein in (A) and (B)

but exists in (C). Bulliform cell (bu); Bundle sheath (bs); Mesophyll (m);

Sclerenchyma (s); Stomatal complex (st); Nodular short-cell (sh-n); Saddle silica-cell

(si-s). Scale bar in (A), (B) and (C) = 200 µm and (a), (b) and (c) = 100 µm.


Figure 10: Changes in internal structure and chlorophyll content during

dehydration and rehydration of Tripogon loliiformis leaves.

Freehand transverse sections of hydrated (A), desiccated (B) and rehydrated (C)

leaves. (D) demonstrates the changes in relative chlorophyll content during

dehydration and rehydration. Dashed line in (A) indicates mestome sheath (me). (B)

White arrow pointing to fragments of vacuole, black arrow pointing to cell wall

folding in mestome sheath and sclerenchyma cell (down), red arrows pointing to

cuticle on abaxial surface (down) and cuticle on bulliform cells (up). Chlorophyll

extraction was performed using N,N-Dimethylformamide of three replicates

(explained at section 4.2.2). Bulliform cell (bu); Bundle sheath (bs); Mesophyll (m);

Mestom sheath (me); Metaxyleme vessel (mv); Sclerenchyma (s); Stomatal complex

(st); Long cell (lo); Phloem (p) and Xylem (x). Scale bar in (A), (B) and (C) = 10

µm.


3.4 DISCUSSION

Leaf structural mechanisms are important aspects of plant environmental stress

tolerance. This study has demonstrated that T. loliiformis has four significant

structural mechanisms that contribute to desiccation tolerance; i) bulliform cells with

large vacuoles, ii) structures on the surface such as trichomes and cuticle, iii) the

unique position of sclerenchymatous tissue and iv) leaf folding. Bulliform cells with

large vacuoles act as water storage units during dehydration (Vecchia et al., 1998).

The presence of epicuticular wax has been associated with dehydration avoidance in

plants (Eglinton and Hamilton, 1967). Macro-hairs and prickle hairs reduce air-

movement over the leaf surface and thus reduce dehydration rate. With the absence

of prickle hairs and macro-hairs on the abaxial surface, air movement increases,

leading to less heat absorbance. The lower heat absorbance property of the smooth

abaxial surface would likely lead to lower leaf temperature. The occurrence of

macro-hairs in T. loliiformis is an adaptation response to extreme environmental

conditions as macro-hairs are absent from leaves grown under high humidity. The

salt glands might dispose the increasing salt concentration during dehydration,

resulting in dehydration prevention ion toxicity.

Dehydration of the bulliform cells (also known as motor cells) is significant

strategy to reduce the surface area through leaf folding in T. loliiformi. In T.

loliiformis bulliform cells have a thin cuticle and cell wall on the adaxial side, which

facilitate water loss and folding. Alvarez et al. (2008) has described the cell walls of

the bulliform cells in two grass species as being rich in pectin. Furthermore, they

describe the bulliform cell walls as being thin and covered by a very thin cuticle.

These characteristics perhaps increase the plasticity of bulliform cell walls which

facilitates the folding. Furthermore, the strategic location of the bulliform cells on

both sides of veins, results in the formation of grooves during dehydration, and these

act as hinges on the sides of veins leading to leaf folding.

The presence/absence and position of sclerenchymatous tissue and other rigid

structures in the leaves of rice have been linked to leaf rolling (Zhang et al., 2009).

Consistent with these observations the changes in the midvein from the apex to the

base of the leaves showed that the absence of sclerenechymatous tissue and other

rigid structures on the adaxial surface facilitate tight leaf folding along the mid-vein

(Figure 9A-C). Where these hard structures appear on the adaxial side of the


midvein, leaf folding is loose (like rolling) as observed at the base of the leaf (Figure

9D-F). Bell and Columbus (2008) demonstrated that leaves of Monanthochloe

littoralis (Poaceae, Chloridoideae) have a V shape similar to folding structure even in

the hydrated state. This plant also lacks the sclerenchymatous tissue and other hard

structures on the adaxial surface of the midvein. Therefore we hypothesise that the

absence of a physical obstacle on the adaxial surface perhaps contributes in the

folding of the leaf at the midvein over adaxial surface.

In T. loliiformis, leaf folding is on the adaxial surface, exposing the abaxial

side to the stressful environment during dehydration. Importantly, the epidermal cells

on the abaxial surface have a thick cuticle (Figures 7,11) thus providing a protective

layer on the exposed surface. Thick-walled sclerenchymatous tissues partially

covered the abaxial surface under epidermis (Figure 7). It is evident that this rigid-

dead tissue shields the internal part of the leaf, protecting the chloroplasts from light

resulting in lowering light induced damage while the leaf is folded.

Cell wall folding reduces tension between the plasma membrane and the cell

wall, preventing the rupture of the cell membrane or cell wall collapse (Vertucci and

Farrant, 1995;Farrant, 2000;Vander Willigen et al., 2004). In Craterostigma wilmsii

cell wall folding resulted in a 78 % reduction in mesophyll cell area following

dehydration to 5 % RWC (Farrant, 2000). Cell wall folding also results in reducing

the overall leaf surface area to minimise light-induced damage. Vander Willigen et

al. (2004) reported that cell wall folding was observed only in thin walled cells such

as mesophyll and epidermis cells of the resurrection grass Eragrostis nindensis.

Although cell wall folding can be seen only in cells with thin wall, this phenomenon

has been associated with the change in chemical composition of the cell walls during

dehydration and it is not an uncontrolled collapse due to dehydration (Moore et al.,

2013).

Increased plasticity of cell walls during dehydration has been attributed to

biochemical changes of the cell wall such as an increase in alpha-expansins,

xyloglucan endotransglucosylases, pectinesterases, pectatelyases and arabinose-rich

polymers (Jones and McQueen-Mason, 2004;Rodriguez et al., 2010;Moore et al.,

2013). A reduction of glucose in the hemicellulose of drying C. wilmsii was reported,

while xyloglucans and unesterified pectins increased (Vicré et al., 1999). Expansins

are other components which increase cell wall flexibility by disrupting the hydrogen


bonds between hemicellulose and cellulose polymers during dehydration (Wu and

Cosgrove, 2000;Jones and McQueen-Mason, 2004;Moore et al., 2008). Different

degrees of cell wall folding have also been reported in different parts of many seeds

(Webb and Arnott, 1982;Roberts, 1986;Woodenberg et al., 2014). Cell wall folding

is important in preserving the structural integrity of the cells during extreme

dehydration and desiccation.

As reported here, fragmentation of the central vacuole has been observed in a

number of desiccation tolerant plants (Gaff et al., 1976;Bartley and Hallam,

1979;Hallam and Luff, 1980;Farrant, 2000;Vander Willigen et al., 2003;Vander

Willigen et al., 2004). Typically, vacuole fragmentation is accompanied by a

concomitant replacement of water with non-aqueous substances and is another

protection mechanism for the plasma membrane during desiccation (Vertucci and

Farrant, 1995;Farrant, 2000). Vacuoles are considered as the major water reservoir of

cells occupying up to 80 % of the cell volume (Alberts et al., 2010). When tissue

dehydrates, the vacuole plays a critical role in cell shrinkage (plasmolysis).

In some resurrection plants such as C. wilmsii, E. nindensis, M. flabellifolius

and X. humilis, the central vacuole is fragmented into multiple smaller vacuoles and

it is suggested that the water in the vacuoles is replaced by non-aqueous substances.

The non-aqueous substances are likely to include osmolytes such as sucrose, proline

and proteins however the exact nature of these compounds is still unknown (Vander

Willigen et al., 2004). Vander Willigen et al. (2004) reported that in Eragrostis

nindensis, these small vacuoles are of different electron density, suggesting the

distribution of metabolites is not uniform in each vacuole. Further,Vander Willigen

et al. (2004) demonstrated that bundle sheath cells with thick cell walls do not

display cell wall folding, however the vacuole in these cells were replaced by

multiple smaller vacuoles.

The fragmentation of the central vacuole and replacement of water with non-

aqueous osmolytes can alleviate mechanical stress by preventing shrinkage as well as

increasing the osmotic pressure (Vander Willigen et al., 2004). While these small

vacuoles constitute nearly 13 % of the total dry mass in dry leaves, they occupy over

30 % of the leaf tissue (Vander Willigen et al., 2004). Fragmentation of the vacuole

also changes the surface to volume ratio (Michaillat and Mayer, 2013), which could

possibly prevent extensive folding of the tonoplast and irreversible fusion during


desiccation. After rehydration the fragmented vacuoles coalesce into a single central

vacuole.

The structural features of T. loliiformis leaves share structural features of plants

from arid areas. These features contribute to desiccation-tolerance by providing the

needed time for developing desiccation-tolerance. Structural changes during

dehydration are to protect the plant against light induced and mechanical damage,

while leaf folding also results in reducing the rate of water loss. All of these features

show that leaf structure plays a critical role in desiccation tolerance in T. loliiformis

and possibly other resurrection plants.


Chapter 4: Physiological Responses of Tripogon loliiformis Leaves During Dehydration and Rehydration 73

Chapter 4: Physiological Responses of

Tripogon loliiformis Leaves

During Dehydration and

Rehydration

4.1 INTRODUCTION

Plants react to dehydration with a range of diverse physiological responses.

Physiological responses are particularly critical for desiccation-tolerance of

resurrection plants. Examples of physiological responses seen in resurrection plants

include changes in photosynthesis, stomata closure, pigmentation and respiration

(e.g. Hsiao, 1973;Farrant, 2000).

One of the main problems plants face during dehydration is oxidative stress

resulting from the accumulation of reactive oxygen species (ROS). This ROS

accumulation is mainly due to the disruption of the photosynthesis electron chain

during dehydration (mostly due to CO2 shortage). Accumulation of ROS can damage

macromolecules (particularly the photosynthetic apparatus), membranes and even

trigger apoptosis in high concentrations. Therefore, the integrity of macromolecules

and membranes is used as a benchmark for determining the level of cellular stress

and ROS damage.

Resurrection plants undergo many physiological changes during dehydration

and rehydration, many of which are designed to minimise water loss and oxidative

damage. Similar to desiccation-sensitive plants, water loss in resurrection plants is

regulated through tight control of stomatal complexes to slow down dehydration.

This provides the time needed for inducing desiccation-tolerance in the vegetative

tissues during dehydration. The accumulation of ROS is minimised mainly through

reducing photosynthesis. Resurrection plants are known to reduce their

photosynthesis faster than desiccation-sensitive plants (Farrant, 2000) perhaps as

they are primed for desiccation. There are a number of mechanisms that plants use to

reduce photosynthesis including, 1) stomatal closure (particularly effective on C3

plants), 2) down-regulation of photosynthetic related gene expression, 3) breaking

down the photosynthetic apparatus (more effective among poikilochlorophyllous

74 Chapter 4: Physiological Responses of Tripogon loliiformis Leaves During Dehydration and

Rehydration

resurrection plants), 4) reducing light-chlorophyll interaction through pigmentation

of leaves and reduction in leaf surface area.

The two main objectives of this chapter were:

1. To characterise the main physiological changes that takes place during

dehydration and rehydration.

2. To determine level of oxidative damage as the result of oxidative stress

through examining the integrity of the photosynthesis apparatus, plasma

membranes and cell viability throughout dehydration, desiccation and

rehydration.


4.2.1 Plant materials and dehydration and rehydration treatment

Plants derived from seeds of a single mother plant were grown for three

months before experiments commenced. Plants were dehydrated by withholding

water until an air-dry state was reached; rehydration occurred for three days. A

minimum of three (n=3) biological replicates were used for each experiment.

4.2.2 Methods

A series of physiological experiments has been conducted in order to monitor

the physiological changes and ROS damage during dehydration and rehydration. The

main physiological changes associated with desiccation tolerance including

chlorophyll integrity and photosynthetic rate were investigated using light

microscopy and photosynthetic rate measurement techniques. The integrity of the

cell membrane and chlorophyll were examined in order to determine the degree of

damage made by ROS on the cells. Cell membrane integrity was determined by

measurement of electrolyte leakage (EL) and confirmed using propidium iodide (PI)

staining. Chlorophyll integrity was tested though measuring chlorophyll content and

chlorophyll fluorescence. Changes in hydration levels have been observed through

RWC and Fresh-Dry weight ratio measurement.


Water content

Relative water content was determined by the standard formula RWC (%) =

(FW-DW)/(TW-DW)*100 (Turner, 1981) [relative water content (RWC); fresh

weight (FW); turgid weight (TW)]. Total leaves from three plants were harvested

(n=3) every day and processed separately. To measure full turgor leaves were placed

in a plastic zip-lock bag with damp filter paper and incubated at 4C overnight. Dry

weight was measured following two days incubation in an oven at 70C.

Quantification of chlorophyll fluorescence and content

To investigate the impact of abiotic stresses and damage to chlorophyll during

dehydration and rehydration, the maximum photochemical quantum yield of

Photosystem II (PSII) (Fv/Fm) and effective photochemical quantum yield of PSII

(ΦPSII) were measured with a portable chlorophyll fluorometer PAM-2500 (Waltz

GmbH, Effeltrich, Germany). Measurements were performed on the third fully

expanded leaf from three leaves (n=3) attached to the plants. Maximum

photochemical efficiency (Fv/Fm) was determined on leaves that were dark-

acclimated for 30 minutes (Kitajima and Butler, 1975). Effective photochemical

quantum yield of PSII in the light adapted state was calculated as the average of the

first five readings using ΦPSII= (Fm′ – Fs)/Fm′ (Genty et al., 1989).

Five plants were used as biological replicates for the measurement of

chlorophyll content; replicates were normalised by dry weight. Extraction of

chlorophyll was done using N,N-Dimethylformamide (Inskeep and Bloom, 1985). To

minimise light and temperature-induced chlorophyll degradation all extracts were

kept on ice and in the dark. Total chlorophyll content were quantified by

spectrophotometric absorbance at 665 and 649 nm and calculated using the modified

formula described in Inskeep and Bloom (1985), total Chl = 17.90A649 + 8.08A665.

For statistical significance, the mean value of three technical replicated

measurements was used, for each biological replicate.

Validation of membrane integrity

Cell membrane integrity was determined by measurement of electrolyte

leakage using a conductivity meter (CM100-2, Reid & Associates CC, Durban,

South Africa). Briefly, three leaves from three biological replicates (plants (n=3))

were cut in 1 cm segments and washed two times in deionized water to remove


Rehydration

surface electrolytes. Residual surface water was removed and the leaf segments were

placed into provided 3.5ml wells that were filled by deionized water. Electrolyte

leakage was measured and calculated according the manufacturer’s instruction.

Propidium iodide (PI) is a membrane impermeable stain and an established

marker of cell membrane integrity. To investigate cell membrane integrity during

dehydration and rehydration three biological and technical replicates were stained

with PI at each hydration point, specifically, hydrated, desiccated and rehydrated.

Samples that had been boiled for 10 minutes in water to disrupt cell membranes

before staining were also included and served as positive staining controls.

Preparation of the PI solution and leaf staining was done as described in Rolny et al.

(2011). Stained leaves were mounted onto slides and examined under an A1Confocal

Microscope (Nikon, Japan). Green images representing nonspecific fluorescence

taken under 488 nm laser were used as contrast to red images as the result of PI

excitation under 561 nm laser.

Quantification of photosynthetic efficiency

Net photosynthesis (A) was measured using a whole Arabidopsis chamber and

a LI-COR 6400-XT InfraRed Gas Analyzer (IRGA) (John Morris Scientific,

Chatswood, NSW, Australia). To minimize erroneous readings due to soil-born CO2

contamination, chamber CO2 levels were maintained above ambient levels. To keep

the CO2 in the chamber above the ambient level, first the level of atmospheric CO2

was measured and then the CO2 supply for the chamber was set to be above that. To

monitor gas exchange through dehydration and rehydration, the same plant was

monitored daily; triplicate plants were assessed. Results are presented as the relative

mean value of photosynthesis rate compared to the photosynthesis rate of the

hydrated plant.

4.3 RESULTS

4.3.1 Leaf water status and pigmentation

Studies have shown that the time taken to dehydrate is vital for the potential of

resurrection plants to resurrect. Plants maintained their leaf RWC for the first few

days before losing approximately 80 % of their RWC within two days (day four and


five). Plants reached an air-dry state (below 10 % RWC) on day six. Upon

rewatering at day six the water level in leaves rose gradually to around 40 % RWC

on day nine (Figure 11). Leaf folding took place at around 65 % RWC and leaves

developed an intense purple pigment intensifying to purple-black on abaxial surface

shortly after leaf folding was completed (Figures 6C,D and 10B). The pigmentation

was quickly lost as plants rehydrated. Plants dehydrated under low light intensity

usually did not develop pigmentation while they are able to resurrect (data not

shown). Desiccation tolerance is usually induced in young and increasingly mature

leaves, while senescing leaves normally do not resurrect. Approximately 30 % of the

total chlorophyll content was lost after dehydration and reconstruction of chlorophyll

after rehydration was slow (Figure 10D).

4.3.2 Chlorophyll a fluorescence

Photosynthesis is particularly sensitive to dehydration stress therefore the

health of the photosystem apparatus, commonly determined by measurement of

Chlorophyll a fluorescence from photosystem II (PSII) is considered a reliable

indicator of ROS levels and damage. To investigate the level of ROS damage during

dehydration and rehydration, the fluorescence of chlorophyll a was observed

throughout dehydration and rehydration; two chlorophyll a fluorescence parameters

were monitored 1) maximum photochemical quantum yield of Photosystem II

(Fv/Fm) (Figure 12A) and 2) effective photochemical quantum yield of PSII (ΦPSII)

(Figure 12B). As expected from a homiochlorophyllous plant the Fv/Fm and ΦPSII

showed similar trends and were almost steady throughout the dehydration and

rehydration period except during day five, six and seven. Severe dehydration during

these three days led to leaf folding. Leaf folding limited the light access to

chlorophylls and made an accurate chlorophyll fluorescence measurement

technically impossible.


Rehydration

Figure 11: Changes in water content during dehydration and rehydration in

Tripogon loliiformis leaves.

(A), changes in relative water content (RWC) and (B), changes in fresh-dry weight

ratio (FW/DW). Rewatering was taken place at day six and the soil was kept moist

for the following three days. Each measurement was done on leaves of three plants.

Figure 12: Changes in chlorophyll fluorescence during dehydration and

rehydration in Tripogon loliiformis leaves.

Triplicate plants were monitored for chlorophyll fluorescence throughout

dehydration and rehydration. Plants were assessed for (A), and (C), maximum

photochemical yield of photosystem II (Fv/Fm), and (B) and (D) for photochemical


quantum yield of PSII (ΦPSII). (C) and (D) demonstrate the average chlorophyll

fluorescence versus relative water content.


Rehydration

4.3.3 Estimation of the membrane integrity

Two types of stress can cause the loss of membrane integrity during

desiccation. Firstly due to increasing tension between plasma membrane and

plasmodesmata connections against the rigid cell wall during severe dehydration

(mechanical damage). Secondly, oxidation and damage of macromolecules in the

membrane by ROS (oxidative stress). Membrane damage leads to the release of

electrolytes from the cells and the rate of electrolyte leakage is linked to the level of

membrane integrity and cell vitality. The level of membrane integrity during

dehydration and rehydration was determined by observation of changes in electrolyte

leakage (EL) and propidium iodide (PI) staining.

Consistent with leaf RWC, the level of electrolyte leakage during the first

three days was stable (Figure 13). As the leaf RWC dropped, the EL rate (ELR)

increased incrementally, by day five, the ELR was 2.5 fold higher than day four

before peaking at a 25 fold increase by day six. Upon rewatering, the ELR was

quickly reduced to approximately 1/3 by day 9 (three days post-watering). A

comparison between changes in RWC and EL shows that the level of EL started to

increase after RWC<90 % and EL rate accelerated sharply by reaching RWC< 60 %.

Two-thirds of the increased EL occurred below 20 % RWC (Figure 13). The

resurrection of T. loliiformis, despite the sharp increase in EL during extreme

dehydration, might suggest that increase in EL might not be the indicator of the

membrane integrity, or the membrane integrity is regained after rehydration. Further

verification of the membrane was needed to observe the integrity of the membrane

during dehydration and rehydration.

To further investigate membrane integrity dehydrating, desiccated and

rehydrated leaves were stained with propidium iodide (PI); hydrated leaves that had

their membranes disrupted by boiling were also included as controls. Propidium

iodide is a membrane impermeable DNA dye that stains the nuclear acid of the cells.

Cells with damaged membrane are permeable to PI and display strong nuclear

staining when excited by fluorescent light. As shown in figure 14, leaves from

hydrated, fully dehydrated and rehydrated plants were impermeable to PI and no

nuclear staining was observed. Contrastingly, PI permeated the boiled controls which

demonstrated strong nuclear staining (Figure 14).


Figure 13: Demonstrating cell membrane integrity in leaves using conductivity

measurement during dehydration and rehydration in Tripogon loliiformis

leaves.

Changes in electrolyte leakage (EL) levels during dehydration and rehydration (A)

and in comparison with changes in relative water content (RWC) (B). Error bars

represent the square root of the standard deviation of the number of samples.


Rehydration

Figure 14: Confocal laser-scanning microscopy of propidium iodide (PI) stained

hydrated, dehydrated (air-dry), rehydrated and control leaves of Tripogon

loliiformis.

Leaves were harvested and stained with PI. Fluorescence was made visible by

confocal microscopy and excitation at 488 nm (left), 561 nm laser (middle) and

merged (right). Green fluorescence images are used as a contrast to red fluorescence

light which represents PI stained area. Stained nuclei can be seen stained as red in

red and merged images of control leaf (bottom). Control leaves were boiled for 10

minutes in order to disrupt the membrane integrity. Scale bar = 50 µm.


4.3.4 Photosynthesis rate

One of the earliest responses of plants to a drop in RWC is the shutdown of

photosynthesis. To further investigate the photosynthetic responses of T. loliiformis

during dehydration we analysed gas exchange and CO2 assimilation using a LI-COR

IRGA. As shown in figure 15, net photosynthesis rate was very sensitive to changes

in hydration levels. At around 90 % RWC, photosynthesis declined by 20 % and

when RWC reached 80 %, photosynthesis was approximately half the level of

photosynthesis before dehydration. There was no measurable photosynthesis at

around 70 % RWC. Photosynthesis was resumed quickly after rehydration (Figure

15).


Rehydration

Figure 15: Changes in photosynthetic rate during dehydration and rehydration

in Tripogon loliiformis.

Error bars represent the square root of the standard deviation of the number of

samples. Three different replicates (n=3) were used for measuring the

photosynthesis.


4.4 DISCUSSION

This study has demonstrated that T. loliiformis has two significant

physiological changes that contribute to desiccation tolerance. These include pigment

accumulation and shutting down the photosynthesis during dehydration.

Furthermore, this study demonstrated that T. loliiformis maintains the integrity of

membrane and much of its photosynthetic apparatus and cells remain viable

throughout dehydration, desiccation and rehydration.

The abaxial epidermis of T. loliiformis is heavily pigmented with anthocyanin

during dehydration (Figures 6D and 10B). This pigmented shield reduces the

excessive radiation injuries and free-radical formation caused by radiation (Bartels et

al., 1992). Dehydration-induced pigmentation is a common feature among

homoiochlorophyllous resurrection plants such as in Craterostigma wilmsii and

Myrothamnus flabellifolius (Sherwin and Farrant, 1998;Farrant, 2000). In T.

loliiformis, this pigmentation appears to be an adaptation response towards high

radiation. Although the pigmentation of the leaves helps with the prevention of

radiation damage during dehydration, the existence of pigmentation seems not

essential in resurrection plants as leaves that are not dehydrated under intense light

and have not produced pigmentation also resurrect (data not shown). Gaff et al.

(2009) also reported similar behaviour in Sporobolus stapfianus when purple

pigmentation was not produced under low light dehydration while plants still

resurrected.

Consistent with observations made by Gaff and McGregor (1979) these results

showed that T. loliiformis maintains most of its chlorophyll during desiccation

(Figures 6,10), which means this plant is a homoiochlorophyllous resurrection plant.

As a homoiochlorophyllous resurrection plant, photosynthesis-induced oxidative

stress is one of the major challenges during desiccation. Reducing light-chlorophyll

interaction through reducing leaf surface area by leaf folding and pigment

accumulation are two strategies that T. loliiformis utilizes to minimise the effect of

light-induced oxidative stress during dehydration. Furthermore, breakdown of 30 %

chlorophyll content could also help with photosynthesis-induced oxidative stress

during desiccation.

While the chlorophyll fluorescence could not be measured during severe

dehydration (when leaves were folded), the chlorophyll fluorescence rate before and

after leaf folding suggested that chlorophyll was protected and remained largely


Rehydration

intact from ROS damage during dehydration (Figure 12). Similar trends for

chlorophyll fluorescence during dehydration and rehydration with a lack of signal

during desiccation were reported in the resurrection plant Haberlea rhodopensis

(Gechev et al., 2013).

Despite increases in conductivity during dehydration, all leaves resurrected,

indicating that cell membranes remain structurally intact during dehydration. Similar

trends in electrolyte leakage (EL) rates were observed by Georgieva et al. (2012)

during dehydration and rehydration in the resurrection plant Haberlea rhodopensis

and increase in EL during desiccation has also been shown in three resurrection

plants Xerophyta humilis, Craterostigma wilmsii and Myrothamnus flabellifolius

(Farrant et al., 2003). This trend of EL in viable tissues of resurrection plants

indicates that the increase in EL during dehydration is not an indicator of membrane

integrity, as loss of membrane integrity is fatal to cells.

Furthermore, Georgieva et al. (2012) also showed that phospholipid

peroxidation did not occur during desiccation, which is an indication of protection

against oxidative damage and membrane integrity. Bajji et al. (2002) suggested the

increase in electrolyte conductivity under osmotic stress in durum wheat is due to an

increase in organic ions and not membrane damage. Rolny et al. (2011) demonstrated

the increase in EL in dark induced senescence in leaves of barley correlated with

ammonium accumulation due to macromolecules degradation; while the cell

membrane integrity was maintained. They suggested the main source of this

ammonium production is from the breakdown of chloroplast which contains up to 70

% protein in the mesophyll cells. The cell membrane integrity during dehydration

and rehydration in T. loliiformis was further proved to remain intact by propidium

iodide staining. A comparison between the staining level of hydrated, dehydrated and

rehydrated tissues with the control boiled sample showed that the cell membrane

remained largely intact (Figure 14).

Gaff and McGregor (1979) observed that during desiccation, non-protein based

nitrogen levels increased significantly in T. loliiformis and decreased during

rehydration while protein-based nitrogen reduced during dehydration and increased

during rehydration. This increase in non-protein based nitrogen and reduction in

protein-based nitrogen is due to the increase in ammonium as the result of protein

breakdown during dehydration, and the increase in protein-based nitrogen and


reduction in non-protein based nitrogen during rehydration is likely due to the flush

of ammonium and the reconstruction of proteins during rehydration.

The increase in EL in T. loliiformis during desiccation is perhaps accompanied

with the breakdown of macromolecules particularly those of photosynthetic

apparatus. In T. loliiformis approximately one third of the chlorophyll is lost during

dehydration. While about two third of the soluble protein is degraded (Gaff and

McGregor, 1979). Therefore, we hypothesize that the main source of this increase in

EL during dehydration is from breakdown of macromolecules. The quick reduction

in EL after rehydration could be due to the electrolytes being flushed by water.

The initial EL increase was at around 90 % RWC, when the plant starts to

dehydrate. This degradation increases as dehydration continues and is accelerated

incrementally from 60 % to the point that 2/3 of the increase in EL took place under

20 % RWC. This perhaps means that the breakdown of macromolecules starts at very

early stages of dehydration as the plant starts to prepare for desiccation. Surprisingly,

the increase in proline concentration was also shown to occur in a very similar

pattern to EL in the resurrection grass E. nindensis (Vander Willigen et al., 2004).

Our speculation is that the accumulation of this amino acid during dehydration could

be in part from breakdown of macromolecules during dehydration.

The initial impact of dehydration on photosynthesis in T. loliiformis started at

the very early stages of dehydration with photosynthesis ceasing at relatively high

levels of hydration (Figure 15). Similar trends have also been reported in a number

of resurrection plants (Tuba et al., 1996b;Tuba et al., 1998;Farrant, 2000). Minor

dehydration triggered reduction in photosynthesis which was perhaps due to the

stomatal closure. In T. loliiformis plants maintained approximately 50 % of their

photosynthetic activity until 80 % RWC while the stomata are closed. Stomatal

closure reduces dehydration rate and provides the time needed for developing

desiccation tolerance and preparing for the desiccation phase. Unlike C3 plants, C4

plants can retain significant levels of photosynthesis while the stomata are closed

(Farquhar and Sharkey, 1982;Flexas and Medrano, 2002). This is due to the C4

photosynthetic mechanism which concentrates the CO2 around Rubisco so that even

the small amount of the CO2 which passes through the cuticle is enough for a

significant level of photosynthesis while the stomata are closed (Boyer et al.,

1997;Ristic and Jenks, 2002).


Rehydration

At around 70 % RWC photosynthesis stops and this coincides with the start of

leaf folding. Photosynthesis resumed quickly upon rehydration to a comparable level

of photosynthesis at same hydration level before dehydration. The quick resumption

of photosynthesis upon rehydration is typical of homoiochlorophyllous DT plants

due to maintaining a significant amount of the photosynthetic apparatus during

dehydration.

Physiological observations related to photosynthesis and electrolyte leakage

(EL) suggest that T. loliiformis is very sensitive to dehydration and starts to prepare

for desiccation at very early stages of dehydration. This quick response to

dehydration could be justifiable based on microhabitats that this plant is commonly

found at i.e. rocky outcrops and shallow rock-pan soils with limited water capacity.

The first response to dehydration is the closing of stomata which results in a

reduction in both photosynthesis as well as transpiration rate.

To conclude, physiological changes protect the plant against oxidative stress

caused by photosynthesis. The photosynthesis is reduced at early stages of

dehydration perhaps through stomatal closure, and later through leaf folding,

pigmentation of the leaves and partial degradation of chlorophyll. Chlorophyll is

protected from oxidative damage during dehydration and rehydration or is repaired

quickly after rehydration. Cell membrane integrity is largely maintained during

dehydration and rehydration. The increase in EL during dehydration is not related to

the loss of membrane integrity and perhaps is related to the release of organic

electrolytes produced during dehydration.


Chapter 5: Protein-protein Interaction Profile of BAG4 Suggests A Possible Role Within Chaperone-mediated

Autophagy 90

Chapter 5: Protein-protein Interaction

Profile of BAG4 Suggests A

Possible Role Within

Chaperone-mediated Autophagy

5.1 INTRODUCTION

Resurrection plants are unique in the ability of their vegetative tissue to tolerate

extreme desiccation but rejuvenate upon addition of water. This trait is reminiscent

of seeds and suggests that Tripogon loliiformis and potentially other resurrection

plants are able to implement effective cytoprotective measures that protect cells and

tissues throughout desiccation. Preliminary research of T. loliiformis shoots

throughout dehydration, desiccation and resurrection performed in the CTCB

suggests that during dehydration and desiccation, T. loliiformis suppresses apoptosis

and senescence pathways whilst promoting pro-survival autophagy pathways

(Williams et al., unpublished). The work performed in this chapter built upon these

interesting findings and was focused on the protein-protein interactions of the

established pro-survival protein, bcl-2-associated athanogene 4 (BAG4). To

investigate whether resurrection plants have evolved unique pro-survival pathways

compared to their desiccation sensitive counterparts, comparative studies of the

protein-protein interactions of BAG4 proteins isolated from Arabidopsis thaliana

(desiccation-sensitive) and T. loliiformis (desiccation-tolerant) were performed.

5.1.1 Programmed cell death

Programmed cell death (PCD) is an innate programme of molecular and

physiological pathways that leads to cell death (Williams and Dickman, 2008).

Programmed cell death is an everyday process and occurs in all multicellular

organisms, including plants, and depending on the context is sometimes triggered in

response to environmental stress. Senescence as a form of cell death occurs in the

vegetative tissues of desiccation-sensitive plants following extreme dehydration

stresses such as salinity and drought where it is thought to be a nutrient recycling

mechanism that benefits the overall plant/organism. Desiccation-tolerant plants avoid


Autophagy 91

senescence even during desiccation. The avoidance of senescence by resurrection

plants during dehydration suggests that these plants might acquire anti-apoptotic

(pro-survival) proteins during that period. The bcl-2-associated athanogene 4

(BAG4) is a molecular chaperone and is one pro-survival molecule that is known to

suppress apoptosis pathways (Doukhanina et al., 2006) and may also be involved in

the repression of senescence.

5.1.2 BAG4

BAG (Bcl-2-associated athanogene) is a family of chaperone regulators which

all share a conserved BAG domain (BD) and has 7 homologues in A. thaliana

(Doukhanina et al., 2006). AtBAG4, a multi-functional protein and remarkably

similar to its mammalian counterpart, is an important regulator of apoptotic-like cell

death in plants. AtBAG4 has been shown to be involved in cell death pathways in

response to a wide range of abiotic stresses as well as being involved in different

growth and given development processes (Doukhanina et al., 2006). Model plants

overexpressing AtBAG4 displayed increased tolerance against a range of abiotic

stresses including UV light, cold, oxidants, and salinity (Doukhanina et al., 2006).

Furthermore, mutant plants displayed a range of growth and developmental changes

including earlier flowering and shorter vegetative and reproductive phases,

producing more branched roots and inflorescences compared with wild-type controls

(Doukhanina et al., 2006). While BAG4 has shown to be involves in a wide range of

developmental stages and abiotic stresses, however the mechanism that this molecule

work is still unknown.

The cDNA of BAG4 from T. loliiformis was isolated and sequenced at Centre

for Tropical Crops and Biocommodities, Queensland University of Technology

(CTCB, QUT) (Williams, unpublished). This gene encodes a 274 aa protein,

however apart from basic sequence analysis the gene is largely uncharacterised. To

date, no BAG family members have been isolated from resurrection plant species.

Due to the observation that T. loliiformis “avoids” PCD even at the desiccation state

(pre-existing T. loliiformis tissues resurrect) and the pro-survival characteristics of

BAG4 proteins, it was feasible to compare protein-protein interaction between

desiccation sensitive and desiccation tolerant plants.

92 Chapter 5: Protein-protein Interaction Profile of BAG4 Suggests A Possible Role Within Chaperone-mediated Autophagy

5.1.3 Protein microarray

The function of a particular protein is largely dictated by its surrounding

proteome and its protein-protein interactions (Waugh, 1954). Traditional methods

such as yeast two-hybrid, in addition to some of the newer fluorescent microscopy

methods enable researchers to study many of these protein-protein interactions.

However, these methods are either laborious, and require, at least to some degree, the

identities of the protein partners to be known or they are also limited in that they only

allow the investigation of single or several interacting proteins at any given time and

thus cannot be used at a large-scale without significant effort. The recent advent of

protein microarrays has emerged as a facile, yet effective means to systematically

analyse potential protein interactions of a given protein of interest. Due to its high

throughput nature, protein microarrays enable the investigation of entire protein-

protein interaction maps (Díez et al., 2012;Sutandy et al., 2013). In essence, protein

microarrays represent a large scale western blot that is performed on a chip that

could have an array of up to thousands of captured proteins. Despite significant

progress in the mammalian field, plant-based protein arrays until recently have not

been available. The recent development of a high-density Arabidopsis protein

microarray enables the study of protein-protein interaction in plants and provides

great potential for the elucidation of plant protein networks. Research reported in this

chapter focused on this investigation and involved the analysis of protein interactions

of AtBAG4 with TlBAG4 to determine whether the T. loliiformis BAG4 protein has

evolved a different suite of protein-protein interaction compared with the

desiccation-sensitive plant, Arabidopsis thaliana. To perform this investigation we

used A. thaliana microarray chips with 5000 proteins fixed on each slide.

The two main objectives of this chapter were:

1. To identify the protein interaction pattern of AtBAG4 and TlBAG4.

2. To investigate the difference between the protein interactions AtBAG4 and

TlBAG4.


Autophagy 93


The molecular studies of this research project used a wide range of molecular

experiments and data analysis method to determine the protein-protein interaction of

BAG4. As was described in section 2.1.3 the technical procedures of the molecular

studies can be divided into five main areas; 1) protein expression, 2) protein

purification, 3) verification, 4) protein microarray and 5) data analysis. An overall

experimental design of molecular work is demonstrated in Figure 16.

Chapter 5: Protein-protein Interaction Profile of BAG4 Suggests A Possible Role Within Chaperone-mediated Autophagy 94

4 sticky tails at 5’ was added and the TAG sequence at 3’ was removed through PCR

Gene was inserted into pENTR vector through

TOPO® Cloning

Genes were inserted into pYL436 vector through LR Cloning

Agrobacterium tumefacients cells were transformed using

electroporation transformation

GUS in Nicotiana benthamiana was expressed through

Agrobacterium-infiltration

Proteins were separated and visualised using SDS Page electrophoresis and

Coomassie blue staining

Protein extraction and purification were done

by IgG sepharose beads

Plasmids was multiplied through transforming and culturing Escherichia coli using heat-shock transformation

Plasmid was extracted from E. coli through

mini prepping

Intactness of plasmid was verified through sequencing

and restriction digestion

Plasmid was multiplied through transforming and culturing E. coli

through heat-shock transformation

Plasmid was extracted from E. coli through

mini prepping

BAG4 was expressed in N. benthamiana through

Agrobacterium-infiltration

Intactness of plasmid was verified through restriction digestion

Existence of the target protein was verified

through Western Blotting

Successful infiltration was verified through

GUS staining

Purified protein was hybridised (probed) against

Arabidopsis protein array slide

The probed microarray was scanned using a Genepix

4300A slide scanner

The protein microarray scanned results was

statistically analysed

Statistical results of protein microarray was analysed by

bioinformatics tools

Figure 16: The experimental design of molecular work procedure.

95 Chapter 5: Protein-protein Interaction Profile of BAG4 Suggests A Possible Role Within Chaperone-

mediated Autophagy

5.2.1 Polymerase chain reaction (PCR)

Reactions were performed in a final volume of 20 µl and included Taq DNA

polymerase (5 units), 400 µM dNTPs 3 mM MgCL2, DNA Dye (Promega), 0.5 µl of

each primer (working concentration of primers was 20 µM), DMSO 10 % (v/v of

final concentration) and autoclaved dH2O. PCR was carried out either in a Peltier

Thermal Cycler – 200 (MJ research - USA) or in a Peltier Thermal Cycler (Gradient

Cycler - Bio-RAD - USA).

The PCR parameters used were;

Denaturation 94˚C 2 min

Denaturation 94˚C 30 sec

Annealing 50˚C 30 sec 30 cycles

Extension 72˚C 70 sec

Final extension 72˚C 10 min

5.2.2 Agarose gel electrophoresis

Agarose gels (1.5 % (w/v)) were made by dissolving agarose (Roche

diagnostics- USA) in 1 X TAE buffer containing 0.5 X SYBR Safe DNA gel stain

(Invitrogen – USA). The EasyCast Mini Gel System was used for casting and

running the gel. Once set, approximately, 12-20 µl of the amplicons including 6 X

bromophenol blue loading were loaded into the gel for electrophoretic separation and

visualisation. Electrophoreses was performed at 120 V for 30 mins unless otherwise

stated. All gels were viewed and photographed using a Syngene Geldocsystem (G-

box and GenSnap version 6.07) (Syngene - UK).

5.2.3 AtBAG4 and TlBAG4

The Arabidopsis thaliana (insert size 840 bp) and Tripogon loliiformis (insert

size 825 bp) BAG4 nucleotide sequences which were extracted from cDNA library

of A. thaliana and T. loliiformis by Dr Brett Williams (CTCB, QUT) and provided to

this project were PCR amplified using primers indicated in Table 5 to remove the

stop codon at the 3’ end and incorporate a 5’ CACC sequence to facilitate directional

Topo cloning.


Table 5: The sequences of primers used for amplification of AtBAG4, TlBAG4

and forward and reverse universal M13 primers used for sequencing.

Primers Sequence 5’ to 3’

AtBAG4 Forward caccatgatgcataattcaaccgaagaatc

AtBAG4 Reverse gtcaaatttctcccaatcttgagttacatt

TlBAG4 Forward caccatgacgggcggcagatcg

TlBAG4 Reverse gtcgaactgctcccagtcagtgttaacttg

Sequencing Forward gtaaaacgacggccag

Sequencing Reverse caggaaacagctatgc

5.2.4 Preparation of bacterial glycerol stock

Bacterial suspension culture was added to equal amount of 80 % glycerol in

Cryovials and snap-frozen before storing at -80˚C.

5.2.5 Extracting plasmid DNA (Mini prep)

Plasmid DNA extraction from Agrobacterium tumefaciens and Escherichia coli

was performed using the standard alkaline lysis method described in Sambrook et al.

(1989). Briefly, a single colony was used for inoculation of 3-4 ml of LB liquid

culture containing appropriate antibody. This liquid was then was incubated on a

shaking rack (225 rpm) at 37˚C overnight for E. coli and at 28 ˚C for 72 h for A.

tumefacient. The liquid culture (2ml for E. coli and 4 ml for A. tumefacient) was then

centrifuged for one minute at 6000 rounds per minute (RPM) at room temperature.

The pellet was resuspended in 100 µl of cold Solution 1 (50 mM glucose, 25 m M

Tris-HCl pH 8.0, 10 mM EDTA). Cells were lysed with the addition 200 µl of

Solution 2 (0.2 M NaOH, 1 % SDS) followed by gentle. Following lysis,

contaminating gDNA and proteins were removed with the addition of 150 µl of

Solution 3 (3M sodium acetate pH 4.7) and 150 µl of CHCL3:IAA (24:1) and

centrifugation at 14000 g for five minutes. The plasmid DNA was precipitated in a

new tube by adding two volumes of 100 % ethanol followed by centrifugation for

five minutes at 14000 g. Excess salts were removed from the plasmid DNA by 70%

ethanol wash. The pellet was air-dried and and dissolved in a final volume of 50 µl

dH2O containing 10 µl RNAse A (10 µg/ml). Plasmids that were used for sequencing


Autophagy 97

were extracted using a Wizard mini prep kit (Promega) according to manufacturer’s

instruction before sending to QUT - Central Analytical Research Facility (CARF)

Molecular Genetics laboratory for sequencing.

5.2.6 Plasmid TOPO® Cloning

TOPO® Cloning was performed according to manufacturers instructions (Life

Technologies) using the primary vector, pENTR and the pYL436 (Gateway-C-TAP)

destination vector that was purchased from the Arabidopsis Resource Center

(ABRC). As described, a 5’ (CACC) was added to both the At and TlBAG sequences

for directional cloning, the stop codon (TAG) was also removed from the 3’

sequence. After PCR the constructs were cloned into pENTR vector using TOPO®

Cloning reaction following the manufacturer’s protocol. The products were used to

transform supplied chemically competent E. coli cells using heat shock as described

in 5.2.7. Escherichia coli cells were cultured on the surface of LB media containing

100 µg/ml kanamycin.

Plasmid DNAs were extracted from positive colonies for further verification on

the target DNA using restriction enzyme digestion and sequencing. Plasmid DNA

was extracted using Wizard mini prep kit (Promega) and the insertion of the target

DNA was verified by using BglII and Sph1 restriction enzymes according to

manufacturer’s instruction and checked for sequence fidelity by sequencing.

After verification, the respective pENTR clones were used in an LR

recombination reaction was used performed, according to manufacturers’

instructions, to transfer the gene to the pYL436 vector (Figure 17). A similar

procedure to that used for the pENTR clones (restriction enzyme digestion and

sequencing) was used to verify the presence of the insert and the fidelity of the target

gene sequence.


Spectinomycin Gentamycin

Figure 17: The position of the gene in pYL436 vector after cloning.

35S promoter; TMV-omega, plant translational enhancer; attL1 and attL2,

recombination sites; BAG gene from Arabidopsis thaliana (AtBAG) or Tripogon

loliiformis (TlBAG); MYC (c-Myc); HIS; C, cleavage site; IgG; Spectinomycin and

Gentomycin coding antibody selection.

pYL436 (12600 bp)

35S AtBAG/TlBAG 9xMYC 6xHIS 3xC 2xIgG attL1 attL2 TMV-omega


Autophagy 99

Once constructed, each expression vector was transformed into Agrobacterium

tumefaciens Agl1 strain cells by electroporation as described in 5.2.7 and selection

on Spectinomycin (25 μg/ml) and Rifampicin (50 μg/ml). Following confirmation of

transformation, a glycerol stock was made from the recombinant Agrobacterium

culture.

5.2.7 Transformation of bacteria

Escherichia coli

The method described by Inoue et al. (1990) was used for transformation of

chemically competent E. coli XL1 Blue cells. Chemically competent cells were

collected from -80 ˚C and left on ice for 10 min. One hundred nanograms of each

plasmid construct was mixed with the bacterial culture in the tube. After 30 min

incubation on ice, tubes were heat shocked at 42 ˚C water for 90 seconds before

transferring on ice for five minutes. The culture was mixed with 500 µl SOC,

incubated at 37 ˚C for 30 min with shaking and spread onto LB media containing the

appropriate antibody overnight.

Agrobacterium tumefaciens

Electro-competent Agrobacterium tumefaciens strain Agl1 were transformed

using an EC100 electroporator (Thermo EC) following the method described by

Dower et al. (1988). Approximately 100 ng of plasmid DNA was mixed with 40 µl

of electro-competent A. tumefaciens cells and transferred to an ice-cold

electroporation cuvette (path length = 2 mm) (Bio-Rad). An electric pulse of 2800 V,

5 msec, 14 kV/cm. 25 µFD was used. The cells were immediately transferred from

the cuvette to a 2 ml tube containing 1 ml SOC and resuscitated by incubation at

28˚C for 2 hours with shaking. Cultures were spread onto LB agar culture containing

spectinomycin (25 μg/ml) and rifampicin (50 μg/ml) for 48-72 hrs at 28 ˚C.

5.2.8 Restriction enzyme digestion of plasmid DNA

Restriction enzyme digestion of plasmid DNA with Mlu1 and DraIII was done

following the manufacturer’s instruction using NEB3 buffer (100mM NaCl, 50mM

Tris-HCl, 10mM, MgCl2, 1mM, DTT, pH 7.9). This was incubated for 1-24 hrs at

37˚C before mixing with DNA Dye (Promega) and electrophoresis.


5.2.9 Agrobacterium-infiltration of Nicotiana tabacum and N. benthamiana

Wild type Nicotiana tabacum and N. benthamiana plants were grown in a

growth chamber under a 16 hrs photoperiod at 25˚C for at least four weeks prior to

infiltration. A two ml of culture of the recombinant Agrobacterium containing the

BAG4 expression cassettes as well as a control consisting of LB media containing

appropriate antibiotic was inoculated by the A. tumefaciens cells (strain Agl1)

containing pYL436 vector with the target genes and (pCAMBIA 2301 vector which

has a GUS gene in it) were used for GUS assay. Cultures were incubated at 28˚C for

48 hrs on a shaker for maximum density. Culture was increased to 10 ml with fresh

LB media containing appropriate antibody and 25 µM acetosyringone (final

concentration) and incubated for 16 hrs.

The culture was centrifuged at 4000 g for 10 min and the pellet was

resuspended and washed twice in 10 ml of infiltration media (10 mM MgSO4.7H2O,

9 mM MES, pH 5.6), before resuspension in 5 ml of infiltration media containing

100 µM acetosyringone. The OD600nm was measured and the culture was diluted to be

an OD600nm between 0.1 to 0.9. Cultures were infiltrated into the abaxial surface of

leaves using 1 ml needle-less syringes. Leaves were retained on the plant for three

days post-infiltration.

5.2.10 Histochemical GUS assay

Infiltrated leaves from N. benthamiana and N. tabacum (pCAMBIA 2301

vector) were used for GUS assay. Agrobacterium-infiltrated leaves were harvested

three days after infiltration and expression level was visualised by adding 5-bromo-4-

chloro-3-indolyl-β-glucuronide (X-gluc) (Progen Biosciences) incubation at 37˚C as

described by Jefferson et al. (1987). Following histochemical staining, chlorophyll

was removed from leaves by incubation in an ethanol:acetic acid solution (3:1)

solution for 48 h.

5.2.11 Protein extraction

The infiltrated region of each leaf was harvested and approximately two grams

were collated, snap-frozen and ground to a fine powder in liquid nitrogen prior to

homogenisation in extraction buffer (100 mM Tris-HCl pH 7.5, 150 mM NaCl, 10%

glycerol, 0.1 % Triton X-100, 1 mM PMSF and 1x complete protease inhibitor

cocktail (Sigma, St. Louise, MO, USA)). Homogenisation was completed by gentle

rotation at 4 ˚C for one hour. Cell debris was removed from the homogenates by


Autophagy 101

centrifugation at 4000 g at 2 ˚C. Further, removal of solids was performed by

centrifugation of the supernatant at 14000 g at 4 ˚C for 20 min. The total protein

extract was incubated with IgG Sepharose 6 Fast Flow beads (Amersham

Biosciences, Uppsala, Sweden) overnight at 4 ˚C with gentle rotation. Recombinant

protein containing the IgG tag was concentrated by centrifugation at 10-12000 rpm

for 3 min at 4 ˚C. The IgG beads and tagged protein were washed 3 times with 1.5 ml

of washing buffer (extraction buffer plus 350 mM NaCl). The recombinant protein

was eluted from the IgG beads through incubation with 12 µl (50 units) of 3C

protease (Precision protease; Amersham Bioshiences) in 500 µl of cleavage buffer

(50 mM Tris HCl pH 7.0, 150 mM NaCl, 1 mM EDTA, 1 mM DTT, 0.1 % Triton)

overnight in cold room with gentle rotation. Supernatant was snap-frozen and stored

at -20 ˚C.

5.2.12 SDS PAGE

SDS-PAGE (sodium dodecyl sulphate polyacrylamide) gel electrophoresis was

used to separate proteins. The protein extract was run on a gel made of 5 % stacking

gel (dH2O 2.916 ml, 1 M Tris pH 6.8 0.5 ml, 10 % SDS 40 µl, 40 % acryl/bis (29:1)

0.5 ml, 10 % ammonium persulphate 40 µl and TEMED 4 µl) and 12 % separation

gel (dH2O 4.135 ml, 1.5 M Tris pH 8.8 2.5 ml, 10 % SDS 100 µl, 40 % acryl/ bis

(29:1) 3.125 ml, 10 % ammonium persulphate 100 µl and TEMED 5 µl). Gels were

prepared using a Mini SDS-PAGE gel apparatus (Bio-Rad) according to the

manufacturers’ protocol. Equal amounts of sample and loading buffer were mixed

followed by a short spin and then heated for 10 minutes at 95 ˚C. A 12 µl aliquot was

loaded to each well and the gels were run for 1 h at 200V. Coomassie blue dye was

used for verification of proteins on the SDS page gel.

5.2.13 Coomassie blue staining

Previously separated proteins were fixed in the SDS gel with the application of

fixing buffer (50 % methanol and 10 % glacial acetic acid) for 1 h. Fixed gels were

stained in Coomassie blue solution (0.1 % Coomassie Brilliant Blue R-250, 50 %

methanol and 10 % glacial acetic acid) for at least 6 hrs with gentle agitation prior to

destaining in a 25 % methanol and 10 % glacial acetic acid solution for one minute

and incubation in 25 % methanol covered by facial tissue until the background

cleared. Images were captured using a Syngene Geldocsystem (G-box and GenSnap


version 6.07) (Syngene - UK) using ‘white background’ option. Gel was stored in

storage solution (5 % glacial acetic acid) until the background colour was removed.

5.2.14 Western blotting

After SDS gel electrophoresis, proteins were transferred to a nitrocellulose

membrane using a Mini SDS-PAGE gel apparatus tank (Bio-Rad). After preparing

the sandwich according to the manufacturers’ protocol, it was placed in the tank

filled with blotting buffer (25 mM Tris, 192 mM glycine, 10 % methanol) and

proteins transferred overnight at 15 V in the cold room. The following day the

apparatus was disassembled and stained with Ponceau Red S (0.1% (x/v) Ponceau S

(Sigma-Aldrich) in 1% (v/v) acetic acid) to verify the transfer of the protein to the

membrane. Ponceau Red S stain was washed off the membrane with tap water.

The membrane was blocked using blocking buffer (TTBS, 5 % (v/w) skim

milk powder) for 1 h with shaking. The blocking solution was discarded and 12 ml

of a diluted 1˚ antibody (mouse (monoclonal) anti-human c-myc unconjugated

antibody) (Invitrogen) solution (1 µg/ml in blocking solution) was added and the

membrane was incubated overnight at room temperature with gentle agitation.

Following incubation, the membrane was washed four times with TTBS (50 mM

Tris, 150 mM NaCl, 0.05 % Tween 20, pH 7.6 (adjusted by HCl)) (10 min each) with

gentle agitation. The membrane was incubated in 2˚ antibody (Goat Anti-Mouse IgG

(H+L) - HRP) (Life Technologies) solution (1 µl of 2˚ antibody in 100 µl blocking

solution, then 9 µl of that solution was mixed in 12 ml of blocking solution)

overnight at room temperature with gentle agitation. Following incubation the

nitrocellulose membrane was washed four times with TTBS as described above. The

chemiluminescent detection assay was performed according to the DIG (Roche)

protocol.

5.2.15 Protein microarray hybridisation (probing) and scanning

A high-density Arabidopsis protein microarray (Dinesh-Kumar Laboratory,

Department of Plant Biology & Genome Centre, University of California, Davis,

USA) with 5000 Arabidopsis ORFs was used to investigate protein-protein

interactions of At-BAG and Tl-BAG proteins. On these protein array slides, each

protein is arrayed in duplicate spots onto glass slides coated with nitrocellulose

polymer (FAST slides; Schleicher & Schuell) and multiple negative (empty) and


Autophagy 103

positive (Cy3 and Cy5) control spots are present. Slides were removed from storage

(-80 ˚C) and placed on ice in an enclosed container for 10 min.

Slides were blocked in blocking solution for 1-2 hours at room temperature and

washed in TBS-1% Tween (50 mM Tris, 150 mM NaCl, 1 % Tween), 3 times, 10

min each wash. Blocked slides were incubated with the eluted proteins (each slide

was treated with one of the proteins) for 1 h while the slides were covered with a

glass slide as hybridization slip. Hybridised slides were washed with TBS-1% Tween

for 10 min, three times. The slides were incubated with primary monoclonal anti-

cMyc IgG (Santa Cruz) diluted 1:2,500 in blocking solution for 1 h while the slides

were covered with a glass slide as hybridization slip. The slides were washed for 10

min in TBS-1% Tween, three times. This was then incubated in the secondary

antibody, Cy5-conjugated anti-mouse IgG (Jackson ImmunoResearch Laboratoires,

Inc.), for 1 hour and washed for 10 min in TBS-1% Tween, three times. Slides were

spin dried by spinning for 1 min at 1000 g inside a 50 ml falcon tube. Slides were

scanned using a Genepix 4300A slide scanner (Axon Instruments).

5.2.16 Statistical analysis

The scanned microarray images were subsequently processed following the

instructions described by Zhou-Da et al. (2008). The report file was generated using

Genepix software to obtain mean, median, and standard deviation of array spots and

background regions. The background intensity of each probe was subtracted from the

corresponding mean intensity and the resulted mean values were used for further

statistical analysis. The average, standard deviation and standard error of the mean

values of the Hsc70 and Ubiquitin spots (each has two replicates) were calculated

and 2* standard error was reduced from the average. All the mean values above this

point were considered as significant for the final analysis.

5.2.17 Bioinformics analysis

Enrichment of gene ontology (GO) terms

GO term enrichment was performed for the protein sets considered to

significantly interact with BAG4 obtained from each protein chip. Each gene set was

enriched using a Fisher’s exact test using the entire annotated transcriptome as a

reference. For statistical significance p-values were corrected according to Benjamini


and Hochberg (1995) and a critical false discovery rate (FDR) q-value of 0.05 was

applied.

Determination of significant protein interactors

The entrez GIs were filtered from the TAIR10 Geneidentifier database that was

downloaded from the Arabidopsis Resource Center (ABRC) website. GIs were

captured using the VLookup function in Excel and were then annotated using the

Kobas online server and Arabidopsis as the reference background. Annotated protein

lists were then used for identification of enriched pathways (KEGGS, Panther and

Biocyc) as well as Gene Ontologies, also performed using KOBAS. For statistical

significance p-values were corrected according to Benjamini and Hochberg and a

critical False Discovery Rate (FDR) q-value of 0.05 was applied.

5.2.18 Quantitative real-time PCR analysis

Superscript III Reverse Transcriptase (Invitrogen) was used to generate cDNA

from 0.8µg of Total RNA using an oligo (dT) (100ρmol) primer. Quantitative PCR

data was generated using a ViiA™7 Real-Time PCR System (Life Technologies) and

the SYBR Green PCR Master Mix kit (Applied Biosystems) according to the

manufacturer’s instructions using 300 µM primer and 1/100 dilution of cDNA and

standard cycling parameters. Gene specific primers for selected genes were designed

using Primer3 software (MIT) and data analysis was conducted using ViiA™ 7

Software v1.2.4 and ExpressionSuite Software v1.0.4 (Life Technologies). The T.

loliiformis homologue of Arabidopsis Actin, Eif and Ubi10 identified from the

annotated transcriptome were used for quantitative normalisation.


Autophagy 105

5.3 RESULTS

Apoptotic and anti-apoptotic proteins play a significant role in growth,

development and response to environmental stresses. Resurrection plants tolerate

desiccation and unlike many other land plants the cells maintain viability suggesting

that they might use anti-apoptotic proteins. An understanding of protein interaction

of these pro-survival proteins might shed light on the mechanism that resurrection

plants use their pro-survival proteins differently compared with desiccation-sensitive

plants. This information might help us in developing stress tolerant crops.

In order to study the protein-protein interaction of the AtBAG and TlBAG,

these proteins were expressed in the leaves of Nicotiana benthamiana through

Agrobacterium-infiltration and purified through IgG sepharose bead purification. The

purified proteins were hybridised against Arabidopsis protein microarray slides and

then scanned using a Genepix slide scanner; results were processed using statistical

and bioinformatics programmes (Figure 16)-(section 5.2.16).

5.3.1 Generation of constructs for Agrobacterium-infiltration

TOPO® Cloning was used to clone the At- and TlBAG4 genes into the target

vector. To facilitate directional cloning and expression of the protein fusion, the

respective BAG4 genes were PCR amplified and site-directed mutagenesis was used to

include four nucleotides (CACC) to the 5’ end of the gene and removal of the stop

codon from the 3’ end.

Constructs were first cloned into the primary “entry” vector (pENTR) by TOPO®

Cloning and then to the final expression vector pYL436 by LR cloning

(Gateway®

cloning). The pYL436 vector contained an IgG epitope and a 3C protease

cleavage site for protein purification as well as nine myc-C and hist tags for detection of

protein-protein interactions. All clones, both entry and destination, were verified by

restriction enzyme digestion and sequencing (Figure 18).


Figure 18: Restriction enzyme digestion of the pENTR and pYL436 for

verification of the insertion of the constructs.

Digestion of the pENTR constructs with Mlu1 resulting in three fragments of

approximately 1473 bp, 1117 bp and 932 bp (left). Restriction enzyme digestion of

pYL436 constructs with DraIII resulting in three fragments of approximately 7632

bp, 3510 bp and 620 bp (right). L, molecular weight standard ladder. White arrow is

pointing at 1000 bp line.

TlBAG4 TlBAG4 AtBAG4 AtBAG4 L L


Autophagy 107

5.3.2 Agrobacterium transformation and infiltration of tobacco and N.

benthamiana

To determine the optimal Agrobacterium-transient expression system both N.

benthamiana and N. tabacum leaves were infiltrated with recombinant

Agrobacterium harbouring the pCAMBIA 2301 vector. A GUS expression vector,

pCAMBIA 2301 is routinely used in molecular biology to quantify transgene

expression levels. Three days post-infiltration, leaves were histochemically assayed

for GUS expression. Since the sole purpose of GUS expression in N. benthamiana

and N. tabacum leaves was for the optimisation of the system and determination of

the best expression host, the levels of GUS expression obtained from pCAMBIA

2301 were estimated based upon visual comparison of the intensity of the blue

staining in the leaves instead of quantitative analysis. GUS assay results were

consistent with previous reports in the lab and revealed that N. benthamiana are more

suitable for expression of the genes using Agrobacterium-infiltration method (Figure

19). Based on these results all transient expression assay were performed in N.

benthamiana.


Figure 19: Determination of the optimal Agrobacterium-transient expression

system.

Leaves of Nicotiana tabacum (left) and Nicotiana benthamiana (right) plants were

infiltrated using Agrobacterium tumefaciens carrying the GUS expression vector

pCAMBIA 2301. Infiltrated leaves were harvested 3 days post-infiltration, GUS

expression was made visible through histochemical GUS assay.

Nicotiana benthamiana Nicotiana tabacum


Autophagy 109

5.3.3 Protein extraction and purification

Once the Agrobacterium-transient expression system was shown to be

working, N. benthamiana leaves were infiltrated using A. tumefaciens carrying the

binary expression vector pYL436 with the inserted target gene constructs. Infiltrated

leaves were harvested three days post-infiltration and total protein was extracted.

AtBAG4 and TlBAG4 constructs were isolated from the total protein extracts using

IgG sepharose beads; the cleavage site on the expressed peptide enables the isolation

of the target constructs from the total protein (Figure 20A). By separating the beads

from the solution and cutting the peptide from the cleavage site, peptides were

successfully released and isolated from the total protein (Figure 20B). The

expression, extraction and isolation were verified by Coomassie blue staining and

Western blotting (Figure 21).

5.3.4 Protein microarray hybridisation (probing) and scanning

Following expression and purification, the AtBAG4 and TlBAG4 proteins

were analysed by Arabidopsis protein chip for protein-protein interactions. Each

high-density Arabidopsis protein microarray slide has 5000 duplicated Arabidopsis

expressed protein spots. As controls, empty (negative) and Cy3 and Cy5 (positive)

spots were also present on the chip.

Independent Arabidopsis chips were hybridised with the AtBAG4 and TlBAG4

proteins. The primary antibody, anti-MYC IgG, detects the MYC peptide which is

originated from the added sequences to the BAG4 proteins (Figure 17B). Anti-MYC

IgG antibody is the target of 2˚ antibody which is an anti-IgG antibody attached to a

Cy5 fluorophore (Cy5-conjugated anti-mouse IgG) (Figure 22). This sandwich was

then detected by a scanner with the presence of Cy5 fluorescence indicative of

protein interaction.


A)

B)

Figure 20: Peptide construct before and after isolation.

A) The peptide expressed in the inoculated Nicotiana benthamiana leaves. B)

Isolated peptide eluted from IgG Sepharose beads after treatment with cleavage

enzyme. BAG protein from Arabidopsis thaliana (AtBAG) or Tripogon loliiformis

(TlBAG); MYC (c-Myc); HIS; C, cleavage site; and IgG.

AtBAG/TlBAG 9xMYC 6xHIS 3xC 2xIgG

AtBAG/TlBAG 9xMYC 6xHIS


Autophagy 111

Figure 21: Verification of expression, extraction and isolation of target proteins

using Coomassie blue staining and Western blotting.

A) Coomassie blue staining of SDS Page gel. B) Western blotting result of the gel

using mouse (monoclonal) anti-human MYC unconjugated antibody as 1˚ antibody

and goat Anti-mouse IgG as 2˚ antibody. At-B/Tl-B, protein solution after overnight

treatment of IgG sepharose beads; At-C/Tl-C, the elution after treatment of IgG

sepharose beads with cleavage enzyme; At-T/Tl-T, total extracted protein from

AtBAG4/TlBAG4 construct treated leaves; L, molecular weight standard ladder;

black arrows pointing at the bands related to cleavage enzyme; red arrows pointing at

the bands related to the constructs.

Tl-T Tl-B Tl-C At-T At-B At-C A

B

L


target protein

(BAG + MYC)

Reporter antibody

(anti-MYC IgG)

capture antibody

(Arabidopsis

protein library)

Sandwich

fluorophore

(Cy5-conjugated

anti-IgG)

A B C D

Figure 22: The procedure of protein microarray chip hybridisation.

Arabidopsis thaliana protein library consisted of 5000 proteins with 2-4 technical replicates as well as positive and negative controls

on each slide. Each slide is treated separately by our target proteins, Tl and AtBAG. BAG protein is attached to a MYC which is

treated by anti-human MYC. This is then treated with Cy5-conjugated anti-mouse IgG. Cy5 is a fluorophore and can be scanned (649

nm absorbance and 670 nm emission wavelength).


mediated Autophagy

5.3.5 Statistical and bioinformatics analysis

For detection of the interacting proteins hybridised slides were scanned using a

GenePix 4300A scanner using 649 nm absorbance and 670 nm emission wavelength

for Cy5 fluorophore. The results of scanning of the hybridised slides showed a large

number of interactions which appeared to have a very similar pattern for both BAG4

proteins (Figure 23). The scanned data was processed using Genepix software and

the report file including mean, median, signal value and standard deviation of array

spots and background regions were maintained. The hybridisation data indicate that

the genes have far more possible interactions than what was suggested/predicted with

the bioinformatics tools. A list of 407 proteins, identified to be above the threshold

significance for AtBAG4 and 671 for TlBAG4 treatment as well as well as compared

AtBAG4 versus TlBAG4 signals can be found in Table 6.

Differential GO term distribution of a given gene set indicates the biological

processes and metabolic functions enriched within that sample. Therefore, once

filtered, each gene set was annotated using KOBAS and subjected to a Fisher’s exact

test for visualisation of differential GO term distribution. The entire annotated

Arabidopsis genome was used as a reference for comparison of the protein sets

within each cluster and enrichment of GO terms specific to that particular sample. A

full overview of the GO categories is presented in Appendix A.

Pathway analysis of the protein interactors can provide further information on

the potential roles that the BAG4 orthologues are playing in A. thaliana and T.

loliiformis. For each protein set, pathway enrichment analysis was calculated using

the online KOBAS server, a summary of the results is provided in Table 7. Briefly,

the interactome for both the Arabidopsis and T. loliiformis proteins were consistent

and largely involved in carbohydrate metabolism pathways.


Figure 23: The scanned Arabidopsis thaliana protein microarray slides

hybridised with AtBAG4 and TlBAG4 constructs.

Hybridised Arabidopsis protein microarray slides were scanned by a GenePix 4300A

scanner using 649 nm absorbance and 670 nm emission wavelength for Cy5

fluorophore.


Table 6: The comparison between the number of spots with significant signal values between AtBAG4 and TlBAG4 hybridised Arabidopsis

thaliana protein results.

The number of spots with significant signal value for AtBAG4 407

The number of spots with significant signal value for TlBAG4 671

The number of proteins shared between AtBAG4 and TlBAG4 405

The number of spots unique to AtBAG4 above the threshold 2

The number of spots unique to TlBAG4 above the threshold 266

Table 7: The pathway enrichment analysis of proteins with significant signal values from AtBAG4 and TlBAG4 microarray protein results.


mediated Autophagy

5.3.6 Generation of constructs for Agrobacterium-infiltration

In order to investigate the possible role of TlBAG4 in T. loliiformis in

desiccation tolerance, the expression level of TlBAG4 during dehydration and

rehydration were observed. Changes in expression of TlBAG4 during dehydration and

rehydration were measured using quantitative reverse transcription polymerase chain

reaction (qRT-PCR). After withholding watering, total mRNA was extracted from T.

loliiformis plants in hydrated, 80 % RWC, 60 % RWC, 40 % RWC, 100 % RWC and

after resuming watering mRNA was extracted after 24 (68 % RWC) and 72 hours (87 %

RWC). Gene expression level remained relatively steady throughout dehydration and

rehydration (Figure 24).


Autophagy 117

Figure 24: qRT- PCR results of TlBAG4 during dehydration and rehydration in

Tripogon loliiformis.

The gene expression level of the TlBAG4 was observed during dehydration and

rehydration using qRT-PCR. Data was generated using three biological replicates

and three technical replicates. All measurements were normalised using three

housekeeping genes, Actin, Eif and Ubi10.

0

5

10

15

20

25

30

35

40

Hyd 80 % RWC 70 % RWC 60 % RWC 40 % RWC 10 % RWC 24 h Reh 72 h Reh


5.4 DISCUSSION

BAG4 has shown to promote abiotic stress-tolerance in transgenic plants

(Doukhanina et al., 2006;Hoang, 2014). To date, no studies have been performed on

a BAG4 homologue isolated from a resurrection plant. Furthermore, our knowledge

about the molecules that BAG4 interacts with is limited to Hsp70 and some

transcription factors. The broad impact and chaperone activity of BAGs suggest that

these proteins might interact with other proteins as well. Using high throughput

protein microarray technology we performed a comparative analysis of the protein-

protein interactions of BAG4, an anti-apoptotic (pro-survival) protein from T.

loliiformis, a resurrection plant, and BAG4 from A. thaliana, a desiccation sensitive

plant. This knowledge can provide us with a better insight into BAG4 performance

and possibly could be used to develop drought-tolerant crops.

The BAG domain shared among BAG proteins interacts with the ATPase

domain of Hsp70 (heat shock protein 70) proteins and regulates the chaperone

activity of them. The BAG domain (BD) of AtBAG4 has exceptional high structural

and charge similarities with human BAG4 (hBAG4) (Figure 25). The AtBAG4

protein is localised in cytoplasm and is known to interact with at least three other

proteins, i) Hsc70, an Arabidopsis heat shock protein; NBR1 (AT4G24690), ii) the

selective autophagy substrate with UBA (ubiquitin-associated) zinc-finger and PB1

domain-containing protein; and iii) a AT hook motif DNA-binding family protein;

and SNF7.1, a vacuolar protein sorting-associated protein 32-2 (Doukhanina et al.,

2006;Consortium, 2011). The interaction of AtBAG4 with Hsc70 was tested

experimentally by pulldown assay (Doukhanina et al., 2006) while yeast 2-hybrid

was used to prove the in vitro interaction of the other proteins with AtBAG4

(Consortium, 2011). Bioinformatic analyses using the predictive proteomics software

(STRING) suggests potential interaction of BAG4 with other proteins including;

AtBAG7; AtBAG5; ATBI1 (BAX inhibitor 1); BIP3, an ATP binding protein;

BZIP28, DNA binding/transcription factor, however these interactions have not been

verified by laboratory based experiments and based on the localisation of some of the

possible proteins interaction seems highly unlikely (Figure 26).


Autophagy 119

A

hBAG4 BD AtBAG4 BD

AtBAG4 BD

hBAG4 BD

B

AtBAG4 BD hBAG4 BD C


Figure 25: Comparison between human BAG4 BAG domain (hBAG4 BD) and

Arabidopsis thaliana BAG4 BAG domain (AtBAG4 BD).

(A) The helices shown as yellow ribbons compose a short antiparallel triple-

helix bundle in both human and Arabidopsis BDs. (B) secondary structure-based

sequence alignment of BDs from AtBAG4 and human BAG4. (C) comparison of the

BDs of AtBAG4 and human BAG4. A three dimensional homology model of the

AtBAG4 BD generated by the SWISS-MODEL web server was compared with the

reported structure of the human BAG4 BD. The residues on the surface of the human

BAG4 BD that are important for Hsc70 interaction are indicated and are compared

with the corresponding residues of the AtBAG4 BD. The water-accessible molecular

surfaces are coloured according to electrostatic potential (upper row) and

hydrophobicity lower row). In the electrostatic potential maps, the blue colour

intensity is proportional to positive charge, white to neutral, and red to negative. In

the hydrophobicity maps, the yellow colour intensity is proportional to the

hydrophobicity of the underlying residues. All molecules are oriented so that the

second and third helices (the Hsc70-binding surface) are facing up front. Modified

from Doukhanina et al. (2006).


Autophagy 121

Figure 26: A schematic illustration of protein-protein interaction of AtBAG4

based on experimental proven and bioinformatics prediction data based on

string-db.org.

The purple lines demonstrate the proven interactions and green lines

demonstrate the interaction predictions.


Although there was a risk that the BAG4 protein from T. loliiformis may not

bind to Arabidopsis proteins on the slide due to the high level of sequence

conservation at the protein level the risk was minimal. The presence of the Hsc70

and ubiquitin-associated (UBA) zinc-finger on the chip and their detection as

potential interactors with both At- and TlBAG4 was increasing the chance and the

AtBAG4 data correlated with experimental data from other labs. These results also

indicated that protein interactors for TlBAG were able to be detected using an

Arabidopsis chip.

Hsc70 is a constitutively expressed chaperone protein involved in facilitating

correct protein folding. It has also been shown to participate in disassembly of

clathrin-coated vesicle protein using the ATPase activity (Newmyer et al.,

2003;NCBI, 2014). The regulation of Hsc70 binding to substrate proteins is by ATP

binding and hydrolysis. The highest affinity of Hsc70 with protein substrates is

formed through the ADP-bound form (Agarraberes and Dice, 2001).

AtBAG4 also interacts with UBA (ubiquitin-associated) zinc-finger which is

associated with the ubiquitin pathway. The Ubiquitin–proteasome pathway (UPP) is

an important protein quality control mechanism as it is the primary cytosolic

proteolytic machinery for the selective degradation of various forms of damaged

proteins, particularly oxidised proteins (Shang and Taylor, 2011). Therefore the

presence of a fully functional UPP system is critical for the cell to cope with

oxidative stress. It comes as no surprise that ubiquitin-mediated protein degradation

pathway is involved in biotic and abiotic stresses (Cui et al., 2012). The role of

ubiquitination in PCD suppression has been shown in Arabidopsis (Marino et al.,

2013) where MYB30 a transcription factor that positively regulates the

hypersensitive cell death responses against biotic stress, is lead to proteasomal

degradation and down-regulation of its transcriptional activity.

Similar to animal cells, external ATP (eATP) (and internal ATP) plays

signalling role in higher plants (Chivasa et al., 2005). While the existence of eATP is

vital to plant cells, stress leads to an increase level of eATP in plants (Jeter et al.,

2004;Chivasa et al., 2005). The increased eATP leads to accumulation of ROS out of

cell membrane in a mechanism that is mediated by NADPH oxidase (Song et al.,

2006) Ca2+

and MAPK (Wong et al., 2007;Demidchik et al., 2009). The role of the

NADPH oxidase induced ROS (Ca2+

mediated) as a critical signalling factor in

regulation of various cell functions such as development, stress response,


Autophagy 123

programmed cell death and in plants is now understood (Pérez-Pérez et al., 2012;Xie

et al., 2014).

ROS stress and nutrient starvation could lead to apoptosis. Autophagy however

reduces the oxidative stress through degradation of damaged proteins which are

whether miss-folded or oxidated and recycles the unwanted proteins during

starvation result in reducing the stress and avoiding apoptosis. The results of protein

microarray hybridisation showed BAG4 interacts with a large number of proteins

which are involved in carbohydrate pathways related to production of ATP and

NADPH which results in activation of the autophagy. Furthermore BAG4 also

interacts with Hsc70 and UBA which are involved in protein degradation in

autophagy. Although the exact role of BAG4 in autophagy survival system is not

clear, the fact that it interacts with numerous members of the autophagy system

highlights its role in this system. The role of BAG4 in autophagy survival pathways

makes it anti-apoptotic protein. The fact that TlBAG4 interacts with more proteins

than AtBAG4 might mean the BAG4 from this resurrection plant might have an

active role in desiccation-tolerance.

To conclude, the protein microarray results suggest that the pro-survival

characteristic of BAG4 are as the result of interaction with proteins involved in

autophagy survival pathway. Although it appears that BAG4 interacts with a large

number of proteins that may also be involved in autophagy procedure, but the exact

role of BAG4 within autophagy pathways remains to be discovered. TlBAG4

appears to have more protein interaction than AtBAG4 which might mean that T.

loliiformis is using this protein differently than A. thaliana. However, the lack of

expression level during dehydration and rehydration suggests that TlBAG4 might not

be directly involved in desiccation tolerance in T. loliiformis. More molecular

experiments needs to be done in order to prove BAG4 interacts with these proteins

and these interactions are directly involved in autophagy. Although there needs to be

other experiments to prove the exact role of BAG4, however the results of high-

density protein microarray provided a valuable platform for future studies.


Chapter 6: General Discussion 125

Chapter 6: General Discussion

6.1 INTRODUCTION

Since their discovery in 1912 biologists have been fascinated with the ability of

resurrection or desiccation-tolerant plants to withstand extreme environments. While

numerous studies have been conducted on these plants, the majority of experiments

have been limited to a small number of species, and these have been mainly from

southern Africa (Table 1). This study used T. loliiformis, an Australian native

resurrection grass as a new resurrection model plant to study desiccation-tolerance

mechanisms. Desiccation-tolerance mechanism has been observed from structural,

physiological and to some extent molecular perspectives using a wide range of

experiments. While T. loliiformis is an ideal model for the study of the unique

tolerance mechanisms of resurrection plants it also has some minor limitations. The

wide range of responses that it demonstrated during dehydration, desiccation and

rehydration, its small size and some of the structural changes during dehydration

were obstacles for some of the observation, particularly physiological experiments.

6.2 STRUCTURAL AND PHYSIOLOGICAL FEATURES AND CHANGES

OF T. LOLIIFORMIS LEAVES ARE TO MINIMISE MECHANICAL

AND OXIDATIVE STRESS DEHYDRATION AND REHYDRATION

Leaf structure plays a critical role during plant stress tolerance, particularly

during water deficit. This study characterised the structural features and changes that

are involved with desiccation-tolerance through leaf structural observation at

different hydration levels. A series of histological methods were optimised in order

to minimise their impact on the tissue in order to investigate the structures of this

plant in the natural state. Tripogon loliiformis possesses structural features that are

common among drought tolerant plants (e.g. thick cuticle, trichomes,

sclerenchymatous tissues and large vacuoles). These structural features facilitate

desiccation tolerance by reducing light damage as well as slowing down dehydration

and providing the much needed time for inducing desiccation tolerance.

126 Chapter 6: General Discussion

As a C4, NAD-ME grass with XyMs+ structure, T. loliiformis has general

physiological features of a drought tolerant plant. C4 plants are generally more

drought tolerant compared to C3 plants. Among C4 plants, NAD-ME type plants are

more tolerant to drought compared with other C4 types, NADP-ME and PEPCK

(Brown, 1999). Furthermore, the XyMS+ structure is a trait characteristically held by

plants grown in arid conditions (Prendergast and Hattersley, 1987).

The water efficiency of C4 plants lie in their ability to concentrate CO2 at the

site of fixation (bundle sheath cells). This increases the photosynthesis-related water

use efficiency of C4 plants by approximately three-fold when compared to their C3

counterparts (Long, 1999). The higher concentration of the CO2 around Rubisco

allows C4 plants to photosynthesize even with low stomatal conductance and hence,

reduces transpirational water loss (Farquhar and Sharkey, 1982;Conley et al., 2001).

This could particularly benefit C4 resurrection plants in two ways. Firstly, it allows

photosynthesis to take place while the stomata are closed, providing more energy as

well as crucial time required for inducing desiccation tolerance mechanisms.

Secondandly, the concentration of the CO2 around Rubisco minimizes the production

of ROS that would otherwise be produced as a result of the disruption of

photosynthesis caused by an incomplete electron transport chain because of low CO2

concentrations, which in turn is due to stomatal closure during dehydration.

Therefore, maintaining the integrity of the photosynthetic apparatus during

desiccation increases the ROS damage more significantly in C3 DT vascular plants

as compared with C4 DT vascular plants. The homoiochlorophyllous resurrection

angiosperms Haberlea rhodopensis and Ramonda serbica switch from C3 to

C4/CAM-type photosynthesis upon dehydration (Markovska et al., 1997;Gechev et

al., 2013). This is likely due to the advantage of C4/CAM photosynthesis over C3

during dehydration.

6.2.1 Bulliform cells appear to play a significant role in gas exchange and leaf

folding during dehydration

It has been suggested that CO2 passing through the cuticle is sufficient for

photosynthesis in C4 plants when the stomatal complexes are closed, because of

concentrations of CO2 around Rubisco (Farquhar and Sharkey, 1982). In T.

loliiformis the cuticle on both surfaces is thick except over the bulliform cells on

adaxial surface, therefore, the main route of CO2 to mesophyll should be through

127

bulliform cells when the stomata are closed. A reduction in wax thickness and an

increase in the number of bulliform cells has been reported on the adaxial leaf

surface of C4 grasses under lower CO2 while C3 grasses of same genus did not show

these changes (Tipping and Murray, 1999). This suggests a role for bulliform cells in

gas exchange in C4 photosynthesis. At around 70 % RWC, photosynthesis stops and

this coincides with the dehydration of bulliform cells. This could be explained as

when bulliform cells dry, the passage of CO2 is closed so this leads to a reduction of

photosynthesis. It appears that bulliform cells have a significant role in gas exchange

in T. loliiformis when the stomatal complexes are closed. This makes these structures

important in shutting down of the photosynthesis when they get dry through

restricting the CO2 absorption. Bulliform cells seem to play various roles during

different phases of dehydration. These cells have large vacuoles which act as a water

reserve. Bulliform cells also play an important role in leaf folding during the

dehydration when they act as pivot to facilitate leaf folding. Shortly after leaf folding

anthocyanin pigments accumulate on the exposed surface (abaxial surface) masking

the interior tissue and further reduces the light penetration. The last stage of the

oxidative protection is the anabolic breakdown of photosynthetic apparatus which

minimises ROS production during the prolonged desiccation state. All these

structural and physiological features and changes in T. loliiformis work together in a

synergy and protect this plant against mechanical and oxidative stresses during

extreme dehydration.

Structural changes during dehydration such as leaf folding, cell wall folding

and vacuole fragmentation result in slowing down of dehydration as well as

preventing the loss of membrane integrity (mechanical damage) and the production

of reactive oxygen spec (ROS) (oxidative stress). The tightness of the observed leaf

folding was associated with the pattern of sclerenchymatous tissue on adaxial

midvein. The absence of sclerenechymatous tissue and other rigid structures on the

adaxial surface facilitate leaf folding along the mid-vein tightly (Figure 8A-C),

while the presence of these rigid structures on the adaxial side of the midvein works

as a physical obstacle which leads to a loose leaf folding (like rolling) at the base of

the leaf (bottom 5 mm) (Figure 8D-F).


6.3 TRIPOGON LOLIIFORMIS MAINTAINS ITS CELLULAR INTEGRITY

DURING DEHYDRATION AND REHYDRATION

The main objectives of physiological studies in T. loliiformis during

dehydration and rehydration were to characterise the physiological changes during

dehydration and rehydration and to observe the level protection of plasma membrane

and photosynthetic apparatus integrity during dehydration, desiccation and

rehydration. The integrity of plasma membrane and photosynthetic apparatus was

used as a benchmark for the level of oxidative stress as the result of ROS

accumulation. The results demonstrated that the physiological changes are mainly to

minimise the oxidative stress and as the result, the integrity of the plasma membrane

and photosynthetic apparatus was largely protected throughout dehydration,

desiccation and rehydration and cells remained viable.

Tripogon loliiformis plants subjected to water deficit maintained a hydration

level of over 80 % relative water content (RWC) before a sudden drop of RWC to

almost air-dry level within a short period of time. Withholding the RWC at high

hydration level is perhaps through the closure of stomata and the sudden dehydration

could be due to soil dryness or through opening of the stomata as has been reported

in the resurrection plant Myrothamnus flabellifolia (Moore et al., 2007). This quick

dehydration could benefit the plant by shortening the time that the plant remains at

intermediate moisture levels which is damaging to the cells because of potential

increased ROS production as a by-product of respiration (Leprince et al., 2000).

Typical of a homoiochlorophyllous resurrection plant, T. loliiformis maintains

much of its chlorophyll during dehydration (approximately 70 %). This makes

photosynthesis-induced oxidative stress a critical perturbation for T. loliiformis

during dehydration. Pigment accumulation and leaf folding during dehydration

minimise the light-chlorophyll interaction. The breakdown of a portion of the

chlorophyll could be due to reducing the stress to a manageable levels via reduced

accumulation of photosynthesis-induced ROS; while recycling of its components

could also be used for producing molecules needed for desiccation-tolerance. The

maintained chlorophyll leads to resumption of photosynthesis quickly upon

rehydration.

The chlorophyll fluorescence results during dehydration demonstrated that

while the majority of the chlorophyll is maintained during dehydration, the integrity

129

of the chlorophyll is also protected. The observed limitation in light access during the

measurement of chlorophyll fluorescence was the result of leaf folding and pigment

accumulation, also suggests that leaf folding and accumulation of pigments

significantly limit chlorophyll irradiation during dehydration. When the leaf is

folded, the vascular tissue and anthocyanin pigmentation shade the chloroplasts

which in return helps reduce light-induced damage and accumulation of ROS during

dehydration.

6.3.1 Increase electrolyte leakage is perhaps due to breakdown of

macromolecules as the result of autophagy during dehydration

Despite the increase in electrolyte leakage (EL) during dehydration the

integrity of the plasma membrane was maintained based on the propidium iodide

staining experiment. The increase in EL during dehydration has also been reported in

other resurrection plants (Farrant, 2000;Georgieva et al., 2012). This increase in

electrolyte leakage in resurrection plants during dehydration is perhaps partly from

organic ions such as ammonium as the result of the breakdown of the

macromolecules as has been observed in some desiccation sensitive plants under

stress (Bajji et al., 2002;Rolny et al., 2011). These macromolecules are most likely

from the photosynthetic apparatus such as chlorophyll. We recommend observation

of the changes in the concentration of the ammonium and chlorophyll content during

various stages of dehydration and rehydration against the changes in EL in this plant.

Apart from organic ions that perhaps are involved in the increased electrolyte

leakage during senescence and desiccation, there might be some other sources of

electrolytes that contribute to the increased EL during such periods. In neutral

solution, ATP is ionized and exists mostly as ATP4−

(Storer and Cornish-Bowden,

1976). The increased levels of eATP as a part of autophagy activation pathway

during senesced and desiccated tissues of desiccation-tolerant plants could play a

significant role in increased electrolyte leakage. Furthermore the accumulation of

ROS produced from NADPH in the extracellular space as the result of stress and/or

starvation can be another source of increased EL during stress and/or senescence,

which does not correspond to loss of membrane integrity. However, to prove this

hypothesis, one needs to measure the concentration of each species of the association

ions present in the mixtures. Perhaps this could be achieved through the method


described in Storer and Cornish-Bowden (1976) for calculation of the true

concentrations of species present in mixtures of associating ions.

6.3.2 Reduction of photosynthesis during early stages of dehydration plays a

key role in activation of autophagy procedure and increase in EL

Photosynthesis in T. loliiformis appeared to be very sensitive in response

dehydration. Stomata closed and photosynthesis ceased at relatively high hydration

levels (Figure 15). The increase in electrolyte leakage (EL) corresponded with the

reduction in photosynthesis starting with the closure of stomata (~90 % RWC) and

acceleration of EL by shutting down of photosynthesis at around 70 % RWC (Figure

13). Coincides in EL could be related with autophagy which starts with the start of

ABA signalling and closure of stomatal complexes. When the photosynthesis stops

this leads to additional starvation which accelerates autophagy degradation of

macromolecules. As explained by Chen et al. (1994) the sugar resources of cell

exhaust quickly and cell starts breaking down the starch using amylase enzymes. The

breakdown of starch has also been observed in a number of resurrection plants during

low hydration levels. Furthermore, the preservation of mitochondrial structural

integrity as well as mitochondrial respiration which has been shown to remain active

observed in a number of resurrection plants until very low hydration level confirms

the respiratory activity (Tuba et al., 1997;Tuba et al., 1998;Farrant, 2000).

Mitochondrial respiration can provide ATP and NAHDPH for autophagy during very

low hydration levels. A further indication of the active autophagy during this period

is the increase in the levels of EL at the very low hydration level (Figure 13). Gaff

and McGregor (1979) have demonstrated that protein based nitrogen decreases

during dehydration, which could be due the amount of protein which is degraded

through autophagy during this time. Autophagy must be playing a critical role in

desiccation-tolerance by degradation and recycling of the damaged and unwanted

proteins (e.g. miss-folded and oxidated) during extreme dehydration.

6.3.3 Plasma membrane is protected from mechanical damage during

dehydration

The integrity of the plasmalemma is protected from mechanical damage during

dehydration through cell wall folding and fragmentation of the vacuole. As described

by Vander Willigen et al. (2004) for the resurrection grass Eragrostis nindensis, cell

131

wall folding was observed in thin walled mesophyll cells while bundle sheath cells

with thick cell walls showed vacuole fragmentation but no cell wall folding; in T.

loliiformis cell wall folding was also observed in mestome sheath cells (Figure 10B).

Fragmented vacuoles in bundle sheath cells were filled with a pale orange-brown

substance (Figure 10B). These components perhaps are the non-aqueous electron

dense substances described by Farrant (2000) and Vander Willigen et al. (2004)

which replaces water in fragmented vacuoles in order to prevent damage to the

plasmalemma during desiccation. While there is some speculation regarding the

nature of these non-aqueous materials, the exact composition of these material

remains unknown. Fragmentation of the vacuole also changes the surface to volume

ratio of vacuoles (Iljin, 1957;Michaillat and Mayer, 2013), possibly preventing the

extensive folding of the tonoplast and the irreversible fusion of tonoplasts during

desiccation. Cell wall folding and vacuole fragmentation have also been reported in

orthodox seeds (Roberts, 1986;Woodenberg et al., 2014).

Chen et al. (1994) have demonstrated an increase in the number of autophagy

vacuoles in the cells undergone autophagy. The observation of fragmented vacuoles

in the desiccated tissues of resurrection plants have been suggested to serve different

purposes and filled with components with different electron density (Farrant,

2000;Vander Willigen et al., 2003). Fragmented vacuoles appeared with different

sizes and colours in T. loliiformis (Figure 10B). We suggest some of these

fragmented vacuoles might be autophagy vacuoles (autophagosomes) for breaking

down and recycling of damaged and unwanted proteins during dehydration.

An increase in the proportion of unsaturated phospholipids in the membrane

might also lead to protection and increased membrane integrity during desiccation

and has been observed for a number of resurrection plants. Hoekstra (2005) reported

a negative correlation between the number of double bonds in polar lipids of

membranes and the longevity of desiccation-tolerant tissues. The phospholipid

content and the polyunsaturated lipid levels in the plasma membrane increased in

desiccation-tolerant attached dried leaves of Sporobolus stapfianus, but decreased in

desiccation-sensitive harvested detached dried leaves (Quartacci et al., 1997;Neale et

al., 2000). Increased levels of unsaturated fatty acids were also observed in all lipid

classes of Boea hygroscopica upon dehydration (Navari‐Izzo et al., 1995). The

reduction in the amount of unsaturated phospholipid in unviable tissues and increase


in viable tissues might be linked to peroxidation of the membrane as the result of

oxidative stress during dehydration.

Unsaturated fats have less tendency to align compared to their saturated fatty

acid counterparts. Therefore an increase in unsaturated phospholipids may result in

increased membrane fluidity (Bartels and Hussain, 2011) or even reducing the

chance of membrane fusion during desiccation. However, the exact role of these

polyunsaturated lipids in desiccation-tolerance is unclear. Moon et al. (1995) and

Gombos et al. (1994) demonstrated unsaturated phospholipid compounds in

thylakoid membrane result in accelerating the recovery of the photosystem II protein

complex from cold photoinhibition. They suggest this phenomena is due to an

increase in membrane unsaturated phospholipids and subsequently increased

membrane fluidity. An increase in plasma membrane fluidity has also been observed

during dehydration in orthodox seeds of cucumber (Amritphale et al., 2000). It is

well known that during senescence a large decrease in cell membrane fluidity results

in membrane deterioration (Borochov et al., 1982;Thompson, 1988). Thus it seems

reasonable to suggest that increased membrane fluidity is a pro-survival strategy for

membrane protection during desiccation in resurrection plants. We further

recommend study of the role of membrane fluidity in desiccation-tolerance

particularly their possible role in preventing membrane fusion. These findings could

have possible implication in developing stress tolerant crop plants.

Based on the changes in chloroplast during dehydration, resurrection plants are

divided in two classes, homoiochlorophyllous and poikilochlorophyllous. Although,

the chloroplasts of both groups undergo changes, the chloroplasts of the

homoiochlorophyllous resurrection plants are significantly less affected than those

present in poikilochlorophyllous resurrection plants. In homoiochlorophyllous

resurrection plants various levels of the chlorophyll are maintained while the rest is

degraded (Gaff, 1981;Sherwin and Farrant, 1998). Some of the changes in

chloroplasts of homoiochlorophyllous resurrection plants during desiccation are in

inner membrane organisation and stacking. Chloroplasts become more round in the

homoiochlorophyllous resurrection plant C. plantagineum (Alamillo and Bartels,

2001). Changes in lipid protein ratio and changes in lipid ratio in thylakoid

membranes have also been reported (Quartacci et al., 1997;Navari-Izzo et al., 2000).

Poikilochlorophyllous resurrection plants on the other hand, lose all chlorophyll and

133

grana and the thylakoid membranes are at least partially degraded or turning to rows

of vesicles when desiccated (Gaff, 1981;Sherwin and Farrant, 1998). Dismantlement

or reduction of photosynthesis apparatus reduces the oxidative stress caused by ROS

accumulation by photosynthesis during water deficit stress.

6.3.4 The possible protection of mitochondria structure during dehydration and

rehydration

While photosynthesis is shutdown early in the dehydration process in

resurrection plants, respiration remains active until very low hydration levels,

providing desiccating tissues with much needed energy (Tuba et al., 1997;Tuba et al.,

1998;Farrant, 2000). The activity of respiration suggests that at least a portion of the

mitochondria remains intact until very low hydration levels. A number of changes

have been observed in mitochondria of desiccated tissues of some desiccation-

tolerant plants. In homoiochlorophyllous C. wilmsii mitochondria retained integrity

and the cristae remained well defined at 5 % RWC (Farrant, 2000). In the

poikilochlorophyllous resurrection sedge Coleochloa setifera, mitochondria were

present in aqueously fixed tissue but contained few cristae (Bartley and Hallam,

1979). In four species of poikilochlorophyllous resurrection plants from the Borya

genus, mitochondria lost some of the peripheral membrane and the number of cristae

were reduced (Gaff, 1981). Meanwhile, after re-greening, mitochondria had more

cristae but there were fewer mitochondria (Gaff, 1981).

These changes to the structure of mitochondria might take place in the near

desiccated state, when respiration is shutdown. It seems there might be a difference

in mitochondria structural changes between homoiochlorophyllous and

poikilochlorophyllous resurrection plants, however, a thorough structural comparison

has not been done in previous studies. Time limitations in this project prevented us

from further investigation the structural changes of chloroplast and mitochondria.

Here we recommend observations on structural changes of chloroplast and

mitochondria to be conducted using cryofixation transmission electron microscopy.

Cryofixation method minimises the structural changes that might occur as the result

of fixation procedure.


6.4 PROTEIN-PROTEIN INTERACTION PROFILE OF BAG4 SUGGEST

A KEY ROLE IN AUTOPHAGY

The fact that desiccation tolerant plants avoid PCD during extreme dehydration

suggests that these plants might utilise pathways that suppress PCD. BAG4 is known

to suppress program cell death (PCD) and is considered an anti-apoptotic protein.

Furthermore, elucidating the molecular mechanisms that underlie the cellular

functions of BAG4 becomes an increasingly important task, particularly in light of

the growing evidence connecting BAG4 overexpression to stress tolerance in model

plants (Doukhanina et al., 2006;Hoang, 2014). The mechanism by which BAG4

facilitates stress tolerance and prevents apoptosis remains to be revealed. Identifying

the proteins that BAG4 interacts with can help to understand the anti-apoptotic

mechanism of this protein. The identification of partner proteins for BAG4 may help

to explain the diverse functions of this protein. This project used high throughput

protein microarray technology to look at the difference between the protein-protein

interactions of the BAG4 proteins from T. loliiformis (TlBAG4) as a desiccation

tolerant plant with A. thaliana (AtBAG4) as a desiccation sensitive plant to see if

they are different or similar.

Our knowledge about plant BAG4 biological pathways is very limited,

however the BAG domain (BD) of AtBAG4 on the residues that are important for

Hsc70 interaction as well as in the charge distribution are remarkably similar to BD

of human BAG4. It is reasonable to assume AtBAG4 interacts with Hsc70 in the

same manner hBAG4 interacts with Hsc70. This means our knowledge about

hBAG4-Hsc4 interactions and regulations could be applicable for AtBAG4-Hsc70.

The N-terminal of the AtBAG4 which interacts with other partners is different from

hBAG4. For understanding the interaction pattern of this site we can use the results

from Arabidopsis protein microarray data.

There are three main different forms of autophagic pathways for degradation of

proteins in lysosomes. Chaperone-mediated autophagy (CMA) is a selective

mechanism for the degradation of soluble cytosolic proteins which is different from

other lysosomal degradation as it is independent from vesicular traffic. In this

pathway proteins are individually recognised by a molecular chaperone attached to

Hsc70 and then this complex interacts with another complex on the membrane of the

lysosome. The protein is unfolded probably by the chaperone complex prior to

135

import inside the lysosome (Salvador et al., 2000;Klionsky, 2005). After entering the

lysosome the protein is degraded and the components are recycled (Majeski and Fred

Dice, 2004;Klionsky, 2005). Plants lack lysosomes and specialised autophagy in

specialised vacuolar (the lysosomal CMV).

There are co-chaperones that interact with Hsc70. These co-chaperones act as

chaperones themselves and regulate the activities of Hsc70 by interacting with its

ATPase domain (Majeski and Fred Dice, 2004). BAG family members including

BAG4 are known to interact with Hsc70 hence positively or negatively regulate

Hsc70 activity in the CMV pathway (Nollen et al., 2000;Alberti et al., 2003). Based

on the data that were generated by the Arabidopsis protein microarray chip as well as

previous studies, AtBAG4 interacts with Hsc70.

A large number of proteins have been identified to have possible interaction

with AtBAG4 based on their signal intensity from the data generated by protein

microarray experiment. Many of these proteins are involved in carbohydrate

pathways (Table 6 and Appendix A).

It seems that the contribution of BAG4 (and perhaps the other BAGs) to stress

tolerance and pro-survival activity is through interaction with proteins involved in

degradation of cytosolic damaged proteins. This is through interaction with proteins

involved in autophagy survival mechanisms. However the impact of BAG4 on the

activation of autophagy through this interaction remains to be revealed. The

degradation of the damaged proteins are perhaps involved with CMV vacuolar and

proteasomal degradation. This project has identified a list of proteins involved in

carbohydrate pathway proteins which perhaps involved in production of NADPH

oxidase induced ROS through production of eATP under stress and starvation

conditions. The list of proteins interacted with BAG4 also has a number of MAPK,

radiation related proteins, and ATP-binding proteins which suggest the plant

autophagy regulation is largely similar to animals. The list also include a number

proteins interacting with transcription factors as well as some metal binding proteins

involved in similar biotic and abiotic stress response as well as development which

their pathways can be investigated.

Despite the shortcoming of this project for providing enough evidence that T.

loliiformis is using the pro-survival protein BAG4 differently in comparison with A.

thaliana, as a desiccation sensitive plant, due to time limitations; however, the study


of protein-protein interaction of BAG4 using high-density protein microarray has

shed light on the possible interaction of this chaperone protein. Furthermore, this

study provided a valuable resource for uncovering the mechanism that this protein

interacts with and pathways it operates. In order to validate the results of protein

microarray observations for protein interaction, further experiments such as co-

immunoprecipitation and/or yeast two hybrid and/or Bimolecular fluorescence

complementation (BiFC) need to be conducted with the candidate proteins.

6.5 SUMMARY

In summary, this work used a combination of structural, physiological and

molecular techniques to investigate the unique structural, physiological and

molecular features that enable T. loliiformis, and potentially other resurrection plants,

to tolerate desiccation. Tripogon loliiformis possesses structural and physiological

characteristics that are common among drought tolerant plants (e.g. thick cuticle,

trichomes and photosynthesis pathway). These features make this plant more

efficient with water usage. Structural changes during dehydration such as leaf

folding, cell wall folding and vacuole fragmentation result in the slowing down of

dehydration as well as preventing the loss of membrane integrity (mechanical

damage) and hence the production of ROS (oxidative stress). Physiological

observations related to photosynthesis and electrolyte leakage (EL) suggest that T.

loliiformis is very sensitive towards dehydration and starts to prepare for desiccation

at very early stages of dehydration. This quick response to dehydration could be

justifiable based on microhabitats that this plant is commonly found at i.e. rocky

outcrops and shallow rock-pan soils with limited water capacity. Bulliforms cells

seem to have a critical role in leaf folding and photosynthesis. The protein results

from protein microarray chip demonstrated that BAG4 has anti-apoptotic properties

due to its interaction with a large number of proteins involved in autophagy

(particularly carbohydrates related pathways leading to autophagy). Furthermore, the

data generated from the protein-protein interaction of BAG4 using high-density

protein microarrays provided a valuable resource for uncovering the

mechanisms/pathways that this protein influences for future research.

Chapter 6: General Discussion 137

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156 Appendices

Appendices

Appendix A: Enriched GO terms for AtBAG4 and TlBAG4

Appendices 157

Enriched GO Terms for Arabidopsis thaliana #Term Input

number Background number

P-Value Corrected P-Value

cytosol 102 1592 8.14E-16 1.70E-12 response to cadmium ion 42 442 4.06E-12 3.23E-09 chloroplast stroma 49 584 4.65E-12 3.23E-09 plastid stroma 49 600 1.12E-11 5.86E-09 response to metal ion 46 559 3.70E-11 1.54E-08 carboxylic acid metabolic process 88 1797 2.27E-08 7.88E-06 glycolytic process 21 184 5.15E-08 1.53E-05 small molecule metabolic process 116 2662 6.72E-08 1.75E-05 oxoacid metabolic process 89 1896 1.11E-07 2.36E-05 organic acid metabolic process 89 1897 1.13E-07 2.36E-05 single-organism carbohydrate catabolic process

32 433 3.01E-07 5.24E-05

pyruvate metabolic process 30 389 3.01E-07 5.24E-05 chloroplast part 58 1092 4.62E-07 7.41E-05 carbohydrate catabolic process 33 469 5.54E-07 8.25E-05 single-organism biosynthetic process

122 2978 6.48E-07 9.00E-05

plastid part 58 1115 8.61E-07 0.000112205 monocarboxylic acid metabolic process

63 1257 9.77E-07 0.000118379

monosaccharide metabolic process 32 460 1.02E-06 0.000118379 pigment metabolic process 26 328 1.11E-06 0.000122006 single-organism catabolic process 61 1215 1.36E-06 0.000141699 transferase activity, transferring one-carbon groups

16 143 2.42E-06 0.000239794

methyltransferase activity 15 129 3.21E-06 0.000304237 small molecule biosynthetic process

63 1317 4.03E-06 0.00034458

monosaccharide biosynthetic process

17 168 4.13E-06 0.00034458

organonitrogen compound biosynthetic process

49 931 4.26E-06 0.00034458

generation of precursor metabolites and energy

32 495 4.30E-06 0.00034458

gluconeogenesis 16 151 4.58E-06 0.000353385 antioxidant activity 14 117 5.00E-06 0.000372363 organic substance catabolic process

69 1504 5.52E-06 0.000396379

hexose biosynthetic process 16 154 5.76E-06 0.000399923 response to inorganic substance 63 1342 7.00E-06 0.000470914 hexose metabolic process 28 416 8.24E-06 0.000536655 glucose metabolic process 27 395 9.13E-06 0.000576393 response to salt stress 40 730 1.29E-05 0.000790313 oxidoreductase activity, acting on peroxide as acceptor

12 96 1.56E-05 0.000917837

organonitrogen compound metabolic process

65 1440 1.61E-05 0.000917837

carbohydrate kinase activity 7 27 1.66E-05 0.000917837

158 Appendices

catabolic process 76 1769 1.67E-05 0.000917837 single-organism metabolic process 176 4993 2.00E-05 0.001068718 alpha-amino acid metabolic process

31 519 2.53E-05 0.00132065

cellular amino acid metabolic process

38 707 3.08E-05 0.001565325

phosphofructokinase activity 5 11 3.17E-05 0.001571327 apoplast 22 315 4.34E-05 0.002101559 response to osmotic stress 40 779 5.03E-05 0.002328853 pigment biosynthetic process 19 252 5.44E-05 0.002441802 kinase activity 53 1152 5.75E-05 0.002441802 carboxylic acid biosynthetic process

48 1009 5.86E-05 0.002441802

organic acid biosynthetic process 48 1009 5.86E-05 0.002441802 peroxidase activity 11 94 6.04E-05 0.002468478 riboflavin metabolic process 4 6 6.88E-05 0.002656528 riboflavin biosynthetic process 4 6 6.88E-05 0.002656528 flavin-containing compound biosynthetic process

4 6 6.88E-05 0.002656528

secondary metabolite biosynthetic process

21 305 7.71E-05 0.002919908

sulfur compound biosynthetic process

27 453 8.36E-05 0.00311019

S-adenosylmethionine-dependent methyltransferase activity

10 81 8.62E-05 0.00315195

phosphotransferase activity, phosphate group as acceptor

5 15 0.000103515 0.003622995

response to cytokinin 18 243 0.000103676 0.003622995 6-phosphofructokinase complex 4 7 0.000106047 0.003622995 6-phosphofructokinase activity 4 7 0.000106047 0.003622995 nucleobase-containing compound kinase activity

6 26 0.000117082 0.003935457

lignan biosynthetic process 5 16 0.000133099 0.00433404 lignan metabolic process 5 16 0.000133099 0.00433404 flavin-containing compound metabolic process

4 8 0.000155953 0.00500009

calmodulin-dependent protein kinase activity

6 31 0.000270338 0.008536124

transferase activity, transferring phosphorus-containing groups

55 1294 0.000282013 0.008771881

protein phosphorylated amino acid binding

4 10 0.000303144 0.009155841

phosphoprotein binding 4 10 0.000303144 0.009155841 chloroplast 125 3560 0.000323895 0.009642814 organonitrogen compound catabolic process

20 320 0.000369104 0.010833989

secondary metabolic process 31 609 0.00037522 0.010860547 O-methyltransferase activity 5 21 0.0003883 0.010976794 phosphotransferase activity, alcohol group as acceptor

30 583 0.000389771 0.010976794

cellular amino acid biosynthetic process

26 477 0.000410442 0.011404827

Appendices 159

response to chemical 124 3552 0.000419375 0.011499717 anthocyanin-containing compound metabolic process

9 82 0.000426408 0.011540695

sulfur compound metabolic process

30 595 0.000533251 0.014067017

single-organism carbohydrate metabolic process

53 1274 0.000571521 0.014888118

cofactor metabolic process 27 517 0.000597176 0.015364373 plastid 125 3626 0.00062277 0.015827484 plastid envelope 28 547 0.00064207 0.016121384 response to molecule of fungal origin

6 38 0.000708643 0.017581105

heme metabolic process 5 25 0.000775116 0.018783034 structural constituent of cytoskeleton

5 25 0.000775116 0.018783034

alpha-amino acid catabolic process 12 151 0.000788018 0.018876195 cellular amino acid catabolic process

12 156 0.001025337 0.024009003

water transport 11 135 0.001061477 0.024308981 fluid transport 11 135 0.001061477 0.024308981 AMP binding 3 6 0.00112556 0.025496384 indole-containing compound metabolic process

11 137 0.001185569 0.026566936

sulfur amino acid biosynthetic process

17 281 0.001356692 0.030078141

phosphorus metabolic process 64 1681 0.001432674 0.031428353 chloroplast envelope 26 525 0.001516549 0.032921744 aminopeptidase activity 3 7 0.001578442 0.0339121 indole-containing compound biosynthetic process

10 123 0.001786529 0.037607341

carbohydrate metabolic process 56 1443 0.001831101 0.038160134 alpha-amino acid biosynthetic process

20 370 0.0019499 0.040233585

indoleacetic acid biosynthetic process

9 104 0.002007645 0.040776781

regulation of cytokinin-activated signaling pathway

3 8 0.00213059 0.040776781

peptide disulfide oxidoreductase activity

3 8 0.00213059 0.040776781

histidine phosphotransfer kinase activity

3 8 0.00213059 0.040776781

glutathione disulfide oxidoreductase activity

3 8 0.00213059 0.040776781

ubiquitin binding 3 8 0.00213059 0.040776781 small conjugating protein binding 3 8 0.00213059 0.040776781 indoleacetic acid metabolic process

9 105 0.002132759 0.040776781

regulation of anthocyanin metabolic process

5 34 0.00260712 0.048948095

protein histidine kinase binding 4 20 0.002643522 0.04918839

160 Appendices

Enriched GO Terms for Tripogon loliiformis #Term Input number Background

number P-Value Corrected

P-Value

cytosol 144 1592 1.27E-15 3.10E-12 response to cadmium ion 60 442 1.32E-13 1.61E-10 response to metal ion 67 559 9.62E-13 7.84E-10 chloroplast stroma 67 584 5.62E-12 3.43E-09 plastid stroma 67 600 1.64E-11 8.01E-09 carboxylic acid metabolic process

129 1797 3.08E-08 1.17E-05

single-organism carbohydrate catabolic process

47 433 3.34E-08 1.17E-05

small molecule metabolic process

174 2662 6.72E-08 2.05E-05

oxoacid metabolic process 132 1896 1.06E-07 2.59E-05 organic acid metabolic process

132 1897 1.09E-07 2.59E-05

carbohydrate catabolic process

48 469 1.21E-07 2.59E-05

single-organism biosynthetic process

189 2978 1.27E-07 2.59E-05

organonitrogen compound biosynthetic process

76 931 2.40E-07 4.52E-05

chloroplast part 85 1092 3.15E-07 5.49E-05 monosaccharide metabolic process

46 460 3.96E-07 6.45E-05

small molecule biosynthetic process

97 1317 5.04E-07 7.69E-05

response to inorganic substance

98 1342 6.27E-07 8.85E-05

glucose metabolic process 41 395 6.87E-07 8.85E-05 plastid part 85 1115 6.89E-07 8.85E-05 organonitrogen compound metabolic process

103 1440 7.95E-07 9.71E-05

hexose metabolic process 42 416 9.78E-07 0.000113755 pyruvate metabolic process 40 389 1.15E-06 0.000127884 response to osmotic stress 64 779 1.65E-06 0.000175088 cellular amino acid metabolic process

59 707 2.61E-06 0.000265751

response to salt stress 60 730 3.35E-06 0.000327259 carboxylic acid biosynthetic process

76 1009 3.80E-06 0.000343534

organic acid biosynthetic process

76 1009 3.80E-06 0.000343534

glycolytic process 24 184 4.43E-06 0.000386909 single-organism catabolic process

87 1215 5.11E-06 0.000429857

generation of precursor metabolites and energy

45 495 5.28E-06 0.000429857

pigment metabolic process 34 328 5.94E-06 0.000460021 monocarboxylic acid 89 1257 6.03E-06 0.000460021

Appendices 161

metabolic process organic substance catabolic process

102 1504 7.41E-06 0.000548562

alpha-amino acid metabolic process

46 519 7.64E-06 0.000548939

kinase activity 82 1152 1.13E-05 0.000768688 sulfur compound biosynthetic process

41 453 1.47E-05 0.00097033

response to chemical 204 3552 1.89E-05 0.00121674 secondary metabolite biosynthetic process

31 305 2.10E-05 0.001292435

cellular amino acid biosynthetic process

42 477 2.12E-05 0.001292435

sulfur compound metabolic process

49 595 2.35E-05 0.001368118

lignan biosynthetic process 7 16 2.41E-05 0.001368118 lignan metabolic process 7 16 2.41E-05 0.001368118 calmodulin-dependent protein kinase activity

9 31 2.48E-05 0.001376733

serine family amino acid metabolic process

26 236 2.73E-05 0.001464273

serine family amino acid biosynthetic process

23 194 2.76E-05 0.001464273

phosphotransferase activity, alcohol group as acceptor

48 583 2.84E-05 0.001477406

transferase activity, transferring one-carbon groups

19 143 3.33E-05 0.001695759

nucleobase-containing compound kinase activity

8 26 4.96E-05 0.002377786

cysteine biosynthetic process 22 191 6.01E-05 0.002821827 catabolic process 111 1769 6.40E-05 0.002925387 gluconeogenesis 19 151 6.47E-05 0.002925387 transferase activity, transferring phosphorus-containing groups

86 1294 6.86E-05 0.003012084

cysteine metabolic process 22 193 6.90E-05 0.003012084 single-organism metabolic process

269 4993 7.06E-05 0.003027542

pigment biosynthetic process 26 252 7.40E-05 0.003115768 hexose biosynthetic process 19 154 8.19E-05 0.003391393 monosaccharide biosynthetic process

20 168 8.44E-05 0.003436393

methyltransferase activity 17 129 9.23E-05 0.003695358 cofactor metabolic process 42 517 0.000112115 0.004417682 alpha-amino acid biosynthetic process

33 370 0.000120787 0.004683852

sulfur amino acid biosynthetic process

27 281 0.000160358 0.006121184

single-organism carbohydrate metabolic process

83 1274 0.000164208 0.006171697

phosphorus metabolic 104 1681 0.000170816 0.006322794

162 Appendices

process apoplast 29 315 0.000184065 0.006711506 response to oxidative stress 42 531 0.000190072 0.00682861 glucose catabolic process 25 255 0.000207097 0.007332432 protein phosphorylated amino acid binding

5 10 0.000228519 0.007862986

phosphoprotein binding 5 10 0.000228519 0.007862986 hexose catabolic process 25 258 0.000243667 0.008154508 monosaccharide catabolic process

25 258 0.000243667 0.008154508

secondary metabolic process 46 609 0.000255507 0.00843518 antioxidant activity 15 117 0.000303599 0.009889246 phosphofructokinase activity 5 11 0.00032133 0.0103225 response to carbohydrate 32 376 0.000325351 0.0103225 carbohydrate kinase activity 7 27 0.000357727 0.011204198 sulfur amino acid metabolic process

28 317 0.000432354 0.013370152

riboflavin metabolic process 4 6 0.000457526 0.013630941 riboflavin biosynthetic process

4 6 0.000457526 0.013630941

flavin-containing compound biosynthetic process

4 6 0.000457526 0.013630941

oxidoreductase activity, acting on peroxide as acceptor

13 96 0.000468122 0.01377858

chloroplast 193 3560 0.000497443 0.014420366 response to cytokinin 23 243 0.000567097 0.015924347 response to fructose 16 139 0.000575451 0.015975297 coenzyme metabolic process 31 374 0.000598671 0.016433177 phosphate-containing compound metabolic process

97 1615 0.000677498 0.018390307

6-phosphofructokinase complex

4 7 0.00069596 0.01848076

6-phosphofructokinase activity

4 7 0.00069596 0.01848076

monocarboxylic acid biosynthetic process

41 552 0.000717889 0.018858091

nucleotide kinase activity 5 14 0.000772992 0.02008958 plastid 194 3626 0.000859575 0.022104661 exopeptidase activity 10 66 0.000954063 0.024210692 protein folding 24 270 0.000979374 0.024210692 phosphotransferase activity, phosphate group as acceptor

5 15 0.000996507 0.024210692

flavin-containing compound metabolic process

4 8 0.001010578 0.024210692

ubiquitin binding 4 8 0.001010578 0.024210692 small conjugating protein binding

4 8 0.001010578 0.024210692

organonitrogen compound catabolic process

27 320 0.001015993 0.024210692

phenylpropanoid metabolic process

18 177 0.001020962 0.024210692

porphyrin-containing 20 208 0.001047881 0.024210692

Appendices 163

compound metabolic process protein metabolic process 157 2867 0.001050484 0.024210692 plastid envelope 40 547 0.001081631 0.024695564 carbohydrate metabolic process

87 1443 0.00110356 0.024772432

tetrapyrrole metabolic process

20 209 0.001105278 0.024772432

water transport 15 135 0.001164295 0.025624977 fluid transport 15 135 0.001164295 0.025624977 peroxidase activity 12 94 0.001209771 0.026317504 S-adenosylmethionine-dependent methyltransferase activity

11 81 0.001217306 0.026317504

structural constituent of cytoskeleton

6 25 0.001329251 0.028026242

indole-containing compound metabolic process

15 137 0.001330759 0.028026242

cis-trans isomerase activity 9 58 0.001456259 0.030407193 calcium ion binding 15 139 0.001516743 0.031137839 chloroplast envelope 38 525 0.001669301 0.033984183 vitamin metabolic process 11 85 0.001724978 0.03477267 isomerase activity 19 202 0.001736498 0.03477267 response to stress 192 3657 0.001980261 0.039014334

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STRUCTURAL PHYSIOLOGICAL AND MOLECULAR CHARACTERISATION OF ...eprints.qut.edu.au/90050/1/Mohammad Reza_Karbaschi_Thesis.pdf · Structural, physiological and molecular characterisation