hung-chi liu - uwa research repository · a systematic in planta survey of 14-3-3 protein...

A Systematic in planta Survey of 14-3-3 Protein

Interactions by Bimolecular Fluorescence

Complementation

This thesis is presented for the degree of Doctor of Philosophy

of The University of Western Australia

Hung-Chi Liu

Biochemistry and Molecular Biology

School of Biomedical, Biomolecular and Chemical Sciences

The University of Western Australia

November 2010

Supervisors: Dr Thomas Martin and Dr Martha Ludwig

Declaration I

Declaration

The work presented in this thesis is my own work except where stated. This work

was carried out in the School of Biomedical, Biomolecular and Chemical Sciences,

Faculty of Life and Physical Sciences, at the University of Western Australia. The

material presented in this thesis has not been presented for any other degree.

Hung-Chi Liu

Nov 2010

Acknowledgement II

Acknowledgement

I would like to express my sincere appreciation to my principal supervisor Dr.

Thomas Martin and my secondary supervisor Dr. Martha Ludwig for their support,

guidance and encouragement on not only scientific research, but everyday life during

the period of my PhD study at the University of Western Australia. Thanks must also

go to all the former and present members in the labs of Dr. Patrick Finnegan, Martha

and Thomas for their shared knowledge and inspiring discussions. I will not forget

the delightful feeling and atmosphere I was able to submerge in when talking about

science during our fortnightly group meetings. Special thanks go to Libby Thomas

for her countless chocolates and handmade cakes that recharged my energy for a lot

of late working hours in the lab.

I thank Dr. Anne Bersoult, Prof. Ian Small, Dr. John Bussell, Dr. Itsara

Pracharoenwattana and Prof. Steven Smith at ARC Centre of Excellence in Plant Energy

Biology, University of Western Australia (UWA, Perth, Australia) for their generosity in

providing plasmids and bacterial materials. My special thanks go to Profs. Claudia

Oecking and Klaus Harter for the BiFC vectors (pSPYNE-35S, pSPYCE-35S, pUC-SPYNE

and pUC-SPYCE), which became the starting material for the BiFC work presented in

this thesis. I am grateful to Mr. John Murphy at the UWA Microscopy Centre for his

technical guidance on the use of confocal microscopes. I would also like to

acknowledge Dr. Roger Hellens and Dr. Phil Mullineaux from the John Innes Centre

(Norwich, UK) for allowing us to use the pGreen vector set, Prof. David Baulcombe at

the Sainsbury Laboratory Plant Biosciences Limited (Norwich, UK) for allowing us to

use the p19 suppressor of gene silencing plasmid and Prof. Mike Jones at Murdoch

University (Perth, Australia) for the kind gift of Nicotiana benthamiana seeds.

I would like to acknowledge UWA for a SIRF/UIS Scholarships and the School of

Biomedical, Biomolecular and Chemical Sciences for the Ad Hoc Scholarship.

Acknowledgement III

Thanks to Mark Ching-Lung Chen for proofreading of my thesis. I am also

thankful to Mark and Tsun-Thai Chai for the many hours we spent together in Perth.

My deepest appreciation goes to my wife, Yin-Chen, for her love and support and

for bringing us four lovely daughters. I am truly grateful to my mom for her endless

support, especially for the help with looking after our triplets. Special thanks also go

to our neighbour aunties in Taiwan, who kindly gave us a big hand by helping us taking

care of our children when I was away from home during the later part of my PhD.

Abstract IV

Abstract

Proteins of the highly conserved eukaryotic 14-3-3 family act as key regulators of

a wide range of biological processes, such as metabolic enzyme regulation, cell cycle

control, ion transport, gene expression, protein assembly and translocation. Such

regulation is achieved by interactions of 14-3-3 homo- and heterodimers with target

proteins. To date, more than one hundred 14-3-3 binding targets have been

identified, mainly through far-Western analysis, immuno-precipitation, yeast

two-hybrid screening, proteomic approaches or in silico predictions. Currently, the

literature proposes two mechanisms that could contribute to 14-3-3 isoform-target

protein specificity. Firstly, it is suggested that the specificity is mediated via

interaction of the 14-3-3 protein’s carboxy-terminal ends with target proteins. On

the other hand, the variability of the amino-terminal ends of 14-3-3 proteins gave rise

to the idea that dimerisation is selective and can therefore contribute to 14-3-3

specificity. Analysing whether 14-3-3s can dimerise freely with each other would

further our understanding of the role the amino termini play during dimerisation and

whether specificity is associated with this.

Bimolecular fluorescence complementation (BiFC) enables the visualisation of

protein-protein interaction in planta. The technique is based on the interaction of

two candidate proteins translationally fused to non-fluorescent fragments of the

yellow fluorescent protein (YFP). Interaction of the candidate proteins leads to

functional YFP reconstitution and fluorescence. In this study, ten out of thirteen

expressed Arabidopsis 14-3-3 isoforms were analysed for dimerisation using a BiFC

approach in transiently transformed Nicotiana benthamiana leaf epidermal cells.

This approach demonstrated that all tested 14-3-3 isoforms were able to freely form

homo- and heterodimers with each other. Although almost all of the 14-3-3 dimers

tested localised to both the cytoplasm and nucleus, some 14-3-3 dimers showed

Abstract V

preferential nuclear localisation. Furthermore, a previously undescribed

endoplasmic reticulum associated distribution of 14-3-3 dimers was observed in

response to induced cell death. Differential cellular distribution patterns of 14-3-3

dimers were also observed in stable transformed Arabidopsis plants expressing 14-3-3

BiFC constructs.

In addition to 14-3-3 dimerisation, this study investigated the interaction of seven

14-3-3 isoforms with a potential target protein, histone deacetylase 2C (HD2C).

Interaction of 14-3-3s with HD2C was exclusively found to be in the nucleus and, most

prominently, in the nucleolus. Interaction with HD2C recruited even those 14-3-3s,

which in dimerisation experiments were classified as mostly cytosolic, to the nucleus

and the nucleolus.

List of abbreviations VI

List of abbreviations

ABA abscisic acid

ABRC Arabidopsis Biological Resource Centre

Ala alanine

AMP adenosine monophosphate

ATP adenosine triphosphate

Arabidopsis Arabidopsis thaliana

BiFC bimolecular fluorescence complementation

bp base pairs

BSA Bovine Serum Albumin

BZR1 brassinazole resistant-1

CaMV Cauliflower mosaic virus

cDNA complementary DNA

DAPI 4’, 6-diamidino-2-phenylindole

DNA deoxyribonucleic acid

E. coli Escherichia coli

ECL enhanced chemiluminescence

EmBP1 Em promoter binding protein 1

ER endoplasmic reticulum

EtBr ethidium bromide

FITC fluorescein isothiocyanate

GFP green fluorescent protein

H2O2 hydrogen peroxide

HA hemagglutinin

HDACs histone deacetylases

HD2C Arabidopsis histone deacetylase 2C

List of abbreviations VII

kDa kilo Dalton

LB medium Luria-Bertani Broth medium

MCS multiple cloning sites

MeJA methyl jasmonate

MES 2-(N-morpholino) ethanesulfonic acid

MQ H2O Milli-Q H2O

MS medium Murashige and Skoog medium

Nicotiana Nicotiana benthamiana

NR nitrate reductase

OD600 optical density at 600 nm

ORF open reading frame

PAGE polyacrylamide gel electrophoresis

PCR polymerase chain reaction

PEG polyethylene glycol

PM plasma membrane

RFP red fluorescent protein

PTGS post-transcriptional gene silencing

rpm revolutions per minute

SDS sodium dodecyl sulfate

Thr threonine

Tris tris (hydroxymethyl) aminomethane

YC (C-YFP) C-terminal fragment of YFP

YFP yellow fluorescent protein

YN (N-YFP) N-terminal fragment of YFP

Table of contents VIII

Table of contents

Declaration................................................................................................................................. I

Acknowledgement ................................................................................................................. II

Abstract ................................................................................................................................... IV

List of abbreviations ............................................................................................................ VI

Table of contents ................................................................................................................ VIII

Chapter 1 General Introduction ........................................................................................ 1

1.1. Introduction ..................................................................................................................... 2

1.2. The early history of 14-3-3 proteins ...................................................................... 2

1.3. Structure of 14-3-3 proteins ..................................................................................... 3

1.4. Modes of 14-3-3 action ................................................................................................ 5

1.5. 14-3-3 proteins in plants ............................................................................................ 7

1.6. Biological processes regulated by 14-3-3 proteins in plants ....................... 7

1.6.1. Metabolism .................................................................................................................................................. 8

1.6.2. Organelle functions ............................................................................................................................... 11

1.7. Redundancy versus specificity: the problem behind the analysis of

14-3-3 function ......................................................................................................... 11

1.8. Bimolecular fluorescence complementation (BiFC): live cell imaging

for protein-protein interaction ........................................................................... 13

1.9. Aims of this project .................................................................................................... 14

Chapter 2 Materials and Methods .................................................................................. 17

2.1. Materials ........................................................................................................................ 18

2.1.1. General Materials ................................................................................................................................... 18

2.1.2. Biological materials .............................................................................................................................. 18

2.1.2.1. Escherichia coli strains used in this study ................................................................................ 18

2.1.2.2. Agrobacterium strains used in this study ................................................................................ 18

2.1.2.3. Plant materials ......................................................................................................................... 18

2.1.2.4. Plasmids and primers ............................................................................................................... 19

2.2. Nucleic acid methods ................................................................................................ 20

2.2.1. Plasmid DNA isolation.......................................................................................................................... 21

2.2.1.1. Plasmid DNA preparation ........................................................................................................ 21

2.2.1.2. Isolation of plasmid DNA using a commercial kit ..................................................................... 22

2.2.2. Restriction enzyme digestion ............................................................................................................ 22

2.2.3. Agarose gel electrophoresis ............................................................................................................... 23

2.2.4. Ligation of DNA fragments into vectors ....................................................................................... 23

2.2.5. Polymerase Chain Reaction (PCR)................................................................................................... 24

Table of contents IX

2.2.6. TOPO TA Cloning® ................................................................................................................................. 24

2.2.7. Sample preparation for automated DNA sequence determination .................................... 25

2.3. Bacterial methods ....................................................................................................... 25

2.3.1. Preparation of competent E. coli cells ............................................................................................ 25

2.3.2. Transformation of E. coli competent cells .................................................................................... 26

2.3.3. Preparation of competent Agrobacterium cells ......................................................................... 26

2.3.4. Transformation of Agrobacterium cells ........................................................................................ 27

2.3.5. Colony PCR ................................................................................................................................................ 27

2.4. Plant methods ............................................................................................................... 28

2.4.1. Plant growth ............................................................................................................................................ 28

2.4.2. Isolation of Arabidopsis mesophyll protoplasts ......................................................................... 29

2.4.3. Polyethylene glycol-mediated transfection of A. thaliana mesophyll protoplasts ....... 30

2.4.4. Agroinfiltration of Arabidopsis and Nicotiana leaves ............................................................. 31

2.4.5. Transformation of Arabidopsis using floral dipping ................................................................ 32

2.4.6. Surface sterilisation of Arabidopsis seeds .................................................................................... 33

2.4.7. Screening Arabidopsis transformants ........................................................................................... 34

2.4.7.1. Screening transformants on plates ..........................................................................................34

2.4.7.2. Screening for transformants using Basta selection on soil-grown plantlets ............................34

2.4.8. Plant genomic DNA isolation ............................................................................................................. 35

2.4.8.1. Genomic DNA isolation using the cetyltrimethyl ammonium bromide (CTAB) method ..........35

2.4.8.2. Genomic DNA isolation using the method of Wang et al. (1993) ............................................36

2.4.9. PCR genotyping of transgenic plants ............................................................................................. 36

2.4.10. Biochemical treatments ....................................................................................................................... 36

2.4.10.1. Hydrogen peroxide treatment .................................................................................................36

2.4.10.2. Methyl jasmonate treatment ..................................................................................................37

2.5. Microscopy and imaging........................................................................................... 38

2.5.1. Fluorescence microscopy .................................................................................................................... 38

2.5.2. Confocal microscopy ............................................................................................................................. 38

2.5.3. Image analysis ......................................................................................................................................... 39

2.5.3.1. Quantification of fluorescence intensity using ImageJ ............................................................39

2.5.3.2. Image deconvolution ...............................................................................................................40

2.6. Protein expression analysis using denaturing gel electrophoresis and

Western blotting ....................................................................................................... 40

Chapter 3 Establishment of a Bimolecular Fluorescence Complementation

Assay System in planta ..................................................................................... 42

3.1. Introduction .................................................................................................................. 43

3.2. Results ............................................................................................................................. 45

3.2.1. Identification of a suitable vector system for a plant based BiFC assay system ............ 45

3.2.2. Construction of pGreen-based BiFC vectors ................................................................................. 49

Table of contents X

3.2.3. Cloning of Arabidopsis 14-3-3 open reading frames ................................................................ 52

3.2.4. Insertion of 14-3-3 ORFs into BiFC binary vectors .................................................................... 57

3.2.5. Generation of an Agrobacterium library carrying pGreen-14-3-3-BiFC constructs ... 61

3.2.6. Establishment of transient BiFC assay systems for 14-3-3 protein interaction

analysis ..................................................................................................................................................... 61

3.2.6.1. Transient BiFC assay in Arabidopsis leaves using agroinfiltration ........................................... 63

3.2.6.2. Transient BiFC assay in Arabidopsis mesophyll protoplasts .................................................... 67

3.2.6.3. Transient BiFC assay in Nicotiana leaves................................................................................. 70

3.2.6.4. Robustness of transient BiFC assays using Nicotiana leaves and reproducibility of results .... 73

3.2.7. Identification of subcellular distribution of 14-3-3-BiFC dimeric complexes ................ 76

3.3. Discussion ...................................................................................................................... 81

3.3.1. Generation of pGreen-based BiFC vectors .................................................................................... 81

3.3.2. The suitability of Arabidopsis mesophyll protoplasts for transient BiFC assays .......... 82

3.3.3. Assessment of BiFC assay using transient expression in Arabidopsis and Nicotiana

leaves via infiltration with Agrobacteria.................................................................................... 83

Chapter 4 A Systematic Survey of Subcellular Dimerisation of Arabidopsis

14-3-3 Isoforms by Bimolecular Fluorescence Complementation ... 87

4.1. Introduction .................................................................................................................. 88

4.1.1. Dimerisation of 14-3-3 proteins ....................................................................................................... 88

4.1.2. Subcellular localisation of 14-3-3 proteins in plants ............................................................... 90

4.1.3. Interaction of 14-3-3s with target proteins ................................................................................. 92

4.2. Results ............................................................................................................................. 95

4.2.1. A database analysis of the subcellular localisation of Arabidopsis 14-3-3 proteins

using the SUBcellular location database for Arabidopsis proteins (SUBA) .................. 95

4.2.2. Arabidopsis 14-3-3 isoforms can form dimers with each other in living plant cells ... 99

4.2.3. Subcellular localisation of 14-3-3-BiFC dimeric complexes ................................................ 101

4.2.3.1. Dual cytosolic/nuclear localisation of 14-3-3 dimers ............................................................. 101

4.2.3.2. 14-3-3 dimers were absent from chloroplast stroma but may be associated with

chloroplastic membranes ................................................................................................ 102

4.2.3.3. Variable nuclear localisation patterns of 14-3-3 dimers were observed ............................... 103

4.2.4. Some 14-3-3 dimers showed preferential subcellular localisations ................................ 105

4.2.5. Discovery of a novel localisation pattern of 14-3-3 dimers ................................................. 110

4.2.6. Investigation into the interaction of 14-3-3 proteins with the histone deacetylase

HD2C ....................................................................................................................................................... 112

4.3. Discussion .................................................................................................................... 116

4.3.1. 14-3-3 isoforms can dimerise freely with each other in vivo .............................................. 116

4.3.2. Subcellular localisation of 14-3-3 dimers ................................................................................... 116

4.3.3. Do nuclear localisation patterns reflect function of 14-3-3 dimers? ............................... 117

Table of contents XI

4.3.4. Is the novel 14-3-3 localisation associated with the ER and with wound induced cell

death? ......................................................................................................................................................120

4.3.5. Seven 14-3-3 isoforms can interact with histone deacetylase 2C .....................................121

4.3.6. 14-3-3-HD2C interactions occurs in the nucleoplasm and nucleolus ..............................122

Chapter 5 Analysis of 14-3-3 Dimerisation in Transgenic Arabidopsis Plants125

5.1. Introduction ............................................................................................................... 126

5.1.1. Arabidopsis 14-3-3 isoforms freely dimerise with each other in a heterologous plant

system .....................................................................................................................................................126

5.1.2. A literature review of 14-3-3 gene expression in Arabidopsis ............................................127

5.2. Results .......................................................................................................................... 129

5.2.1. A large number of endogenous 14-3-3s are co-expressed in Arabidopsis......................129

5.2.2. Generation of transgenic Arabidopsis plants constitutively expressing 14-3-3-split

YFP fusion proteins ............................................................................................................................131

5.2.3. Phenotypic changes were observed in some of the T1 transformants ............................134

5.2.4. Genetic analysis of T2 populations of 14-3-3-split YFP transformants ..........................134

5.2.5. Generation of “double” transgenic 14-3-3-YN and 14-3-3-YC Arabidopsis plants for

BiFC analysis of 14-3-3 dimerisation in a whole plant context ........................................137

5.2.5.1. Selection of BiFC expressing, double transgenic Arabidopsis plants .....................................138

5.2.5.2. Distribution of 14-3-3 dimers in double transgenic Arabidopsis plants as observed by BiFC 139

5.2.5.3. Genetic analysis of F2 offspring of double transgenic BiFC Arabidopsis plants .....................141

5.2.6. Distribution of 14-3-3 BiFC dimers in double transgenic Arabidopsis plants ..............143

5.2.6.1. Distribution of 14-3-3 dimers in roots of transgenic seedlings ..............................................143

5.2.6.2. Subcellular distribution of 14-3-3 dimers in cotyledon guard cells ........................................145

5.2.6.3. Subcellular distribution of 14-3-3 dimers in epidermal cells of rosette leaves ......................148

5.2.6.4. Distribution of 14-3-3 epsilon-YN/mu-YC dimer in flower tissues .........................................150

5.3. Discussion ................................................................................................................... 150

5.3.1. BiFC fluorescence in transgenic plants may reflect endogenous 14-3-3 dimerisations150

5.3.2. Visualising 14-3-3 dimerisations in a whole plant context .................................................152

5.3.3. Subcellular distribution of 14-3-3 dimer was somewhat dimer-dependent in some

tissues ......................................................................................................................................................154

Chapter 6 Cell Death Induced Aggregation and ER Association of 14-3-3

Dimers ................................................................................................................. 156

6.1. Introduction ............................................................................................................... 157

6.2. Results .......................................................................................................................... 159

6.2.1. Co-localisation of 14-3-3 dimers with fluorescent ER and Golgi markers .....................159

6.2.2. Chemical cell death inducers and wounding trigger changes of subcellular

localisation of 14-3-3 dimers in transgenic Arabidopsis ....................................................167

6.2.2.1. The subcellular distribution of 14-3-3 dimers changes in response to wounding in cotyledon

epidermal cells ................................................................................................................169

Table of contents XII

6.2.2.2. Subcellular distribution of 14-3-3 dimers in mesophyll protoplasts derived from double

transgenic Arabidopsis plants ......................................................................................... 171

6.2.2.3. Hydrogen peroxide treatment induces the novel subcellular localisation of 14-3-3 dimers . 173

6.2.2.4. Methyl jasmonate treatment induces the novel subcellular localisation of 14-3-3 dimers .. 174

6.3. Discussion .................................................................................................................... 180

6.3.1. The potential association with the ER upon cell death induction and wounding

suggest a role of 14-3-3s in ER mediated apoptosis ............................................................. 180

Chapter 7 General Discussion ...................................................................................... 183

7.1. Conclusion and discussion .................................................................................... 184

7.1.1. Analysis of 14-3-3 isoform specificity using bimolecular fluorescence

complementation ............................................................................................................................... 184

7.1.1.1. Isoform specificity in 14-3-3 dimerisation ............................................................................. 184

7.1.1.2. Isoform specificity with respect to 14-3-3 isoform-target interaction .................................. 185

7.1.1.3. Differential subcellular distribution of 14-3-3 dimers may contribute to specificity ............. 186

7.1.2. Advantages and limitations of bimolecular fluorescence complementation for the

analysis of 14-3-3 interactions ..................................................................................................... 188

7.1.3. Interpretation of the BiFC outcomes of 14-3-3 interactions in cells ................................ 191

7.2. Future perspectives ................................................................................................. 193

References ........................................................................................................................... 196

Appendices .......................................................................................................................... 213

Appendix I: Media and solutions ..................................................................................... 214

Appendix II: Supplementary figures and tables ........................................................ 218

Chapter 1. General introduction 1

Chapter 1

General Introduction


1.1. Introduction

Plants use many strategies to deal with environmental challenges because of

their sessile lifestyles. Challenges such as nutrient availability, light conditions,

abiotic and biotic stresses greatly affect resource acquisition and allocation and thus

the productivity of plants. In response to these challenges, plants integrate various

sensing and signalling mechanisms and exhibit a variety of responses to adapt to

surrounding conditions. Accumulating evidence has shown that signalling pathways

leading to these responses share steps or cross-talk to each other (Genoud and

Métraux, 1999). Understanding these signalling mechanisms is key to improving

crop productivity.

Reversible protein phosphorylation is a major mechanism for signal

transduction at the cellular level (Cohen, 2002). Phosphorylation can regulate

enzyme activity, cellular distribution of proteins, and assembly and disassembly of

protein complexes (Bridges and Moorhead, 2005). However, phosphorylation and

dephosphorylation of proteins, in many cases, are not sufficient for completion of

signalling cascades and even for the cross-talk of diverse signalling cues (Chevalier et

al., 2009). It appears that in many cases proteins that can interact with

phosphorylated signalling components play a role as intermediates to integrate

multiple signalling pathways. A class of proteins known as 14-3-3 proteins is one

group of these intermediate regulators (Bridges and Moorhead, 2005).

1.2. The early history of 14-3-3 proteins

14-3-3 proteins were discovered in 1967 by Moore and Perez during the

classification of bovine brain proteins (Moore and Perez, 1967). The name is based

on the fraction numbers of these proteins during isolation by diethylaminoethyl

cellulose chromatography and starch gel electrophoresis. After their discovery, the


role of 14-3-3s remained elusive for twenty years when it was shown that 14-3-3

proteins act as activators of tyrosine and tryptophan hydroxylases involved in

neurotransmitter biosynthesis (Ichimura et al., 1987). In 1992, 14-3-3 proteins from

plants were first described (Brandt et al., 1992; de Vetten et al., 1992; Hirsch et al.,

1992; Lu et al., 1992), drawing the attention to the role of “brain proteins” in plants.

Since then, it has been shown that 14-3-3s function as key regulators in many

biological processes in both plants and animals by physically interacting with target

proteins in a phosphorylation-dependent manner. It was further shown that 14-3-3

proteins act as protein dimers (Jones et al., 1995; Liu et al., 1995; Xiao et al., 1995).

To date, more than 300 14-3-3 interacting proteins have been identified or

proposed, including proteins involved in a wide range of biological processes such as

metabolic enzyme regulation, cell cycle control, ion transport, gene expression,

protein assembly and translocation (reviewed in Ferl et al., 2002; Comparot et al.,

2003; Roberts, 2003; MacKintosh, 2004).

1.3. Structure of 14-3-3 proteins

14-3-3 proteins, from all eukaryotes, are encoded by small gene families

comprising of two genes in the model yeast Saccharomyces cerevisiae to thirteen

expressed genes in the model plant Arabidopsis thaliana (van Heusden et al., 1996;

Rosenquist et al., 2001; Chevalier et al., 2009). The 14-3-3 family is a highly

conserved protein family, with high amino acid sequence similarities even between

animal and plant homologs, as shown by a phylogenetic analysis involving 153 14-3-3

protein sequences across 48 species (Rosenquist et al., 2000; Ferl et al., 2002). The

overall amino acid identity between the expressed Arabidopsis isoforms is 50% but

can be as high as 92% between two isoforms (Wu et al., 1997a). In general, large

blocks of conserved amino acids are found in the central region of the proteins, with


both the N- and C-terminal ends of 14-3-3s showing greater variation (Chung et al.,

1999; Ferl et al., 2002).

From the highly conserved protein sequences, it is anticipated that the

individual 14-3-3 isoforms share similar protein structures. Indeed, X-ray crystal

structure analyses of seven human and one plant isoforms confirmed that 14-3-3s

form very similar dimeric structures (Liu et al., 1995; Xiao et al., 1995; Rittinger et al.,

1999; Würtele et al., 2003; Gardino et al., 2006). As shown for the human 14-3-3

zeta dimer in Fig. 1-1, each 14-3-3 monomer consists of nine -helices arranged in an

anti-parallel fashion forming an L-shaped structure. The monomer interior consists

of four -helices containing charged and polar amino acids that form a concave

amphipathic groove, which interacts with mostly phosphorylated target peptides.

The less conserved C-terminal domains of 14-3-3 proteins are assumed to play a role

in target protein recognition and to contribute to isoform specificity (Chung et al.,

1999). In addition to the role in substrate recognition, the C-terminus of some

isoforms may also have an autoinhibitory role (Shen et al., 2003).

14-3-3 proteins are known to exist as homodimers and heterodimers (Jones et

al., 1995; Liu et al., 1995; Xiao et al., 1995; Chaudhri et al., 2003). The N-terminal

domains of two 14-3-3 monomers form the dimerisation interface. Structural

analysis of the human 14-3-3 zeta dimer suggests that dimer formation requires the

interaction of the N-terminal helix H1 of one monomer with helices H3 and H4 of the

other monomer (Liu et al., 1995). It is possible that the N-terminal sequence

variability determines which dimers are formed under physiological conditions and

hence it may contribute to specificity (Chung et al., 1999; Aitken et al., 2002);

however, this hypothesis lacks experimental support.


Fig. 1-1. Crystal structure of a 14-3-3 dimer bound to mode-1 target peptides.

Crystal structure analysis of a 14-3-3 dimer bound to mode-1 target peptides shows that the

central structure of the dimer forms a basket-like structure composed of anti-parallel

alpha-helices. Each monomer of the dimer creates a groove that forms a target binding

interface. The conservation of 14-3-3 sequences across large evolutionary distances allows

this structure to serve as an appropriate model for the fundamental structure of all 14-3-3s.

The extreme N- and C-termini are unresolved in this analysis as well as in similar 14-3-3 crystal

structure analysis. This image was created using Cn3D and PDB:1QJB. Reprinted from Physiol.

Plant. 120, Ferl, R. J., “14-3-3 proteins: Regulation of signal-induced events”, pp. 173-178,

copyright 2004, with permission from John Wiley and Sons.

1.4. Modes of 14-3-3 action

Plant and animal 14-3-3 proteins recognise the same set of conserved

phosphopeptide binding motifs, RSXpS/pTXP (mode-1) or RXXXpS/pTXP (mode-2)

(Yaffe et al., 1997; Bridges and Moorhead, 2005). However, binding motifs differing

from these two typical motifs were reported for some 14-3-3 target proteins, such as

the YpTV (Y: tyrosine, pT: phosphothreonine and V: valine) motif at the extreme

C-terminal end of the plant plasma membrane H+-ATPase (Würtele et al., 2003). In a

few examples, 14-3-3 proteins appear to bind to unphosphorylated target proteins (Fu

et al., 2000). The variety of binding motifs and the lack of recognisable motifs in a


great number of putative client proteins indicate that our current understanding of

14-3-3 target protein recognition is still incomplete.

14-3-3 proteins have no intrinsic enzyme activity and their biological function is

only revealed by their interaction with target proteins (MacKintosh, 2004). These

functions can be classified into several modes of 14-3-3 action (Roberts, 2003):

(a) Interaction of 14-3-3s with a client protein, where the client is an enzyme, can alter

the activity of the client protein by changing its specific activity or its turnover rate

(reviewed in Huber et al., 1996; Bunney et al., 2002; Comparot et al., 2003; Lillo et

al., 2004). Well-characterised examples are nitrate reductase (NR) and the plant

plasma membrane H+-ATPase. Binding to 14-3-3 inactivates NR activity whilst it

activates H+-ATPase activity (Oecking et al., 1994; Bachmann et al., 1996a; Huber et

al., 1996; Moorhead et al., 1996; Jahn et al., 1997; Oecking and Hagemann, 1999).

(b) 14-3-3 dimers can act as a scaffold, bringing two client proteins together or

stabilising oligomerisation of a target protein. Examples of this scenario are the

Bcr-Raf protein complex in mammalian cells (Braselmann and McCormick, 1995)

and the VP1-EmBP1 transcription factor complex regulating abscisic acid

(ABA)-responsive gene expression in plants (Schultz et al., 1998). In both cases

14-3-3 dimers bridge and connect the client proteins within the complexes. In

addition, 14-3-3 binding results in H+-ATPase hexamerisation (Kanczewska et al.,

2005; Ottmann et al., 2007), which also supports the adapter mode of 14-3-3

action.

(c) Binding of 14-3-3s to multiple interaction sites within a single target protein may

lead to conformational changes of the target, as is the case in 14-3-3 interaction

with Raf-1 (Tzivion et al., 1998; Yaffe, 2002).

(d) Facilitate import of nuclear-encoded chloroplast proteins into chloroplasts is

stimulated by 14-3-3s bound to the transit peptides of the precursor proteins (May

and Soll, 2000; Sehnke et al., 2000).


(e) 14-3-3s regulate nucleo-cytoplasmic shuttling of some nuclear proteins, such as

the plant transcription factor REPRESSION OF SHOOT GROWTH (RSG) and

mammalian histone deacetylases (HDACs). Here, interaction with 14-3-3s

sequesters RSG or HDACs in the cytosol (Grozinger and Schreiber, 2000; Eckardt,

2001; Igarashi et al., 2001).

(f) In animal cells, a model has been proposed that suggests that interaction with

14-3-3 proteins facilitates transport of correctly assembled multimeric

channels/receptors from the endoplasmic reticulum (ER) to the plasma membrane

(Nufer and Hauri, 2003).

1.5. 14-3-3 proteins in plants

Multigene families encoding 14-3-3s were found in both monocotyledonous (e.g.

barley and rice) and dicotyledonous (e.g. tomato and Arabidopsis) plants. At least

five 14-3-3 encoding genes, named Hv14-3-3A to Hv14-3-3E, are found in barley

(Schoonheim et al., 2007b). In rice, all of the eight identified 14-3-3 genes in the

genome are expressed and are given the names OsGF14a to OsGF14h (Chen et al.,

2006; Yao et al., 2007). In tomato, twelve 14-3-3 genes (TFT1 to TFT12; Roberts,

2003; Xu and Shi, 2006) were identified, whilst Arabidopsis has fifteen 14-3-3 genes,

named GRF1 to GRF15 of which thirteen are expressed (Rosenquist et al., 2001;

Sehnke et al., 2006). To differentiate the Arabidopsis isoforms, individual members

are designated by Greek names and letters (Table 1-1).

1.6. Biological processes regulated by 14-3-3 proteins in

plants

Plant 14-3-3 proteins interact with a wide range of target proteins and are


Table 1-1. The Arabidopsis 14-3-3 family.

Gene name Greek name

of protein Greek letter AGI number*

GRF1 Chi χ At4g09000

GRF2 Omega ω At1g78300

GRF3 Psi ψ At5g38480

GRF4 Phi φ At1g35160

GRF5 Upsilon υ At5g16050

GRF6 Lambda λ At5g10450

GRF7 Nu ν At3g02520

GRF8 Kappa κ At5g65430

GRF9 Mu μ At2g42590

GRF10 Epsilon ε At1g22300

GRF11 Omicron ο At1g34760

GRF12 Iota ι At1g26480

GRF13 Pi π At1g78220

GRF14 - - At1g22290

GRF15 - - At2g10450

* AGI number: Arabidopsis thaliana Gene Index. - Protein name has not been given since the corresponding gene is

proposed to be a pseudogene.

assumed to regulate or impact on such diverse biological functions as plant

metabolism, gene expression, protein translocation into organelles, signal

transduction, hormone biosynthesis and cell cycle regulation (reviewed in Ferl et al.,

2002; Comparot et al., 2003; Roberts, 2003; MacKintosh, 2004). In this section,

some of the well-known processes that are regulated by 14-3-3 proteins in plants are

highlighted.

1.6.1. Metabolism

A large number of metabolic enzymes, most of which are involved in nitrogen

and carbohydrate metabolism, generation of proton gradients and energy


metabolism, are regulated by 14-3-3 proteins (Huber et al., 2002; Comparot et al.,

2003; Chevalier et al., 2009). One of the best-studied roles of plant 14-3-3s is the

inhibition of NR activity upon transition from light to dark, which is regulated by a

two-step process (Bachmann et al., 1996a; Moorhead et al., 1996). Nitrate

reductase catalyses the reduction of nitrate to nitrite, the first step of nitrogen

assimilation in plants (Huber et al., 1996; Campbell, 1999). The first step in the

inhibition process involves the phosphorylation of NR at a serine residue within a

conserved peptide motif by a sucrose non-fermenting related kinase (SnRK) and/or a

calcium-dependent protein kinase (CDPK; Su et al., 1996; Douglas et al., 1998; Sugden

et al., 1999). Subsequently, the phosphorylated NR is bound by 14-3-3 dimers

leading to inactivation of the enzyme (Mackintosh et al., 1995; Bachmann et al.,

1996a). The binding of 14-3-3s to phosphorylated NR is dependent on millimolar

concentrations of a divalent cation, such as Ca2+ or Mg2+ (Lu et al., 1994; Athwal et al.,

1998a), and loop 8 of 14-3-3 proteins is involved in the cation binding (Athwal and

Huber, 2002). Further evidence that 14-3-3 proteins play a role in the inactivation of

NR was obtained by the mutation of the 14-3-3 binding site of tobacco NR, which

resulted in the constitutive activation of the enzyme (Lillo et al., 2003). In addition,

binding of 14-3-3s leads to degradation of the enzyme by an unknown mechanism

(Sehnke and Ferl, 2000). The interaction of 14-3-3s and NR can be disrupted by

adenosine monophosphate (AMP), leading to activation of the enzyme (Athwal et al.,

1998b). This and a putative AMP binding site present in some 14-3-3s suggest that

14-3-3 activity may be under metabolic control (Athwal et al., 1998b).

In addition to NR, 14-3-3 proteins also bind to both the cytosolic and

chloroplastic isoforms of glutamine synthetase (GS1 and GS2, respectively;

Finnemann and Schjoerring, 2000; Riedel et al., 2001). Darkness and high ATP/AMP

ratios lead to phosphorylation of GS1 and subsequent binding of 14-3-3s, causing high


GS1 activity (Finnemann and Schjoerring, 2000). For GS2, only the phosphorylated

and 14-3-3 bound form of the isoform is catalytically active (Riedel et al., 2001).

Thus, regulation of nitrogen metabolism is strongly controlled by the interaction of

14-3-3 proteins with nitrogen assimilating enzymes.

Numerous enzymes involved in carbohydrate metabolism are found to

associate with 14-3-3s, such as sucrose-phosphate synthase (SPS; Toroser et al., 1998),

trehalose-6-phosphate synthase (TPS; Moorhead et al., 1999), starch synthase III

(Sehnke et al., 2001), glyceraldehyde-3-phosphate dehydrogenase (GAPN; Bustos and

Iglesias, 2003) and fructose-2,6-biphosphatase (Kulma et al., 2004). Direct evidence

for the regulation of carbohydrate metabolism by 14-3-3s came from antisense

approaches, which showed that a reduction of the granule-associated Arabidopsis

14-3-3 isoforms epsilon and mu by antisense approaches led to increased starch

accumulation in leaves (Sehnke et al., 2001).

Another well-studied example is the regulation of the plant plasma membrane

(PM) H+-ATPase by 14-3-3s. The primary role of PM H+-ATPase is the generation of a

proton gradient across the PM to provide an energy source for transport of nutrients

into cells, phloem loading and opening and closure of stomata (Palmgren, 2001).

Binding of 14-3-3 proteins to the phosphorylated, C-terminal autoinhibitory domain

of the PM H+-ATPase activates the pump (Jahn et al., 1997; Baunsgaard et al., 1998;

Olsson et al., 1998; Oecking and Hagemann, 1999; Svennelid et al., 1999; Maudoux et

al., 2000). Addition of the fungal phytotoxin fusicoccin to the 14-3-3-H+-ATPase

complex leads to irreversible activation of the pump that results in consistent opening

of stomata and thereby wilting of the leaf (de Boer, 1997; Jahn et al., 1997; Oecking

et al., 1997).


1.6.2. Organelle functions

Together with heat shock protein 70, 14-3-3s facilitate translocation of

phosphorylated chloroplast precursor proteins into wheat chloroplasts (May and Soll,

2000). In addition, 14-3-3s bind to a thylakoid-targeted precursor protein, the

Arabidopsis photosystem I N-subunit (At pPS1-N) as shown using yeast two-hybrid

assays (Sehnke et al., 2000). It was further demonstrated that four Arabidopsis

14-3-3 isoforms, epsilon, mu nu, and upsilon, localised to the chloroplast stroma

despite having no import signals of their own (Sehnke et al., 2000). In another study,

14-3-3 proteins were found in the matrix protein fraction of isolated mitochondria

using immunoblotting analyses (Bunney et al., 2001). Potential target proteins for

14-3-3 action in mitochondria, as well as chloroplasts, were the organelle specific ATP

synthases. Binding of the ATP synthases to 14-3-3s were in a phosphorylation-

dependent manner and down-regulated the activity of the ATP synthases (Bunney et

al., 2001). The mechanism of 14-3-3 translocation into organelles is currently unclear.

Co-translocation with bound target proteins across the organelle membranes is a

commonly proposed mechanism for divergent subcellular localisations of 14-3-3s

since they contain no apparent signal sequences (Paul et al., 2005).

1.7. Redundancy versus specificity: the problem behind the

analysis of 14-3-3 function

The high conservation of amino acids in the internal groove suggests that all

14-3-3 proteins have the general ability to interact with the same subset of target

proteins (Aitken et al., 2002; MacKintosh, 2004). This is likely to account for the

ability of 14-3-3s to complement for each other, even across species. For example,

several 14-3-3 isoforms from Arabidopsis could complement a lethal yeast mutant in

which the two 14-3-3 genes BMH1 and BMH2 were non-functional (van Heusden et


al., 1996). Furthermore, several in vitro and in vivo studies showed that many 14-3-3

isoforms can interact with the same target protein (Bachmann et al., 1996b;

Kanamaru et al., 1999; Rosenquist et al., 2000; Schoonheim et al., 2007a; Sullivan et

al., 2009). For example, it was shown that twelve recombinant Arabidopsis 14-3-3

isoforms were able to bind to the leaf cell PM H+-ATPase by using far-Western blotting

analysis (Alsterfjord et al., 2004). These results can be taken as indication of a high

degree of functional redundancy within 14-3-3 proteins.

On the other hand, differential target protein binding affinities between 14-3-3

isoforms were detected in a number of studies. For instance, it was shown that

seven of the Arabidopsis 14-3-3 isoforms were able to bind to NR in a yeast

two-hybrid assay (Kanamaru et al., 1999). The strongest affinity to NR was observed

for 14-3-3 omega, followed by and in the order phi, kappa, lambda, psi and chi, with

upsilon showing a very poor affinity. Alternative approaches, however, have shown

somewhat conflicting results. Bachmann et al. (1996) used synthetic phospho-NR

peptides to study the interaction between 14-3-3 isoforms and NR and also showed

the strongest binding to NR was by 14-3-3 omega. However, in contrast to the work

of Kanamaru et al. (1999), Bachmann and co-workers found that the 14-3-3 isoforms

chi and upsilon interact strongly with NR whilst phi showed only weak interaction in

their test system (Bachmann et al., 1996b). As shown by Athwal et al. (2000) these

differences in experimental outcome strongly depend on the parameters used in the

test system, for example, the binding affinities of 14-3-3s to NR changed dramatically

when the pH or the divalent cation concentration was altered (Athwal et al., 2000).

The discrepant results discussed above reveal a profound weakness of

interaction studies that are based on in vitro experiments or take place in

non-homologous in vivo systems such as the yeast two-hybrid analysis. Taking into

account that 14-3-3 proteins have a high degree of sequence similarity, especially in


the substrate binding internal groove, a degree of redundancy in the function of

14-3-3s is not surprising. Furthermore, those test systems do not take into account

that 14-3-3 genes are differentially expressed as shown for a large number of plant

isoforms (Daugherty et al., 1996; Roberts and Bowles, 1999; Rosenquist et al., 2001).

The subcellular localisation of 14-3-3s and target proteins may also determine

whether an interaction takes place as 14-3-3 proteins can be localised in the cytosol,

nucleus, chloroplasts and mitochondria (reviewed in Comparot el al., 2003).

Additionally, a 14-3-3 isoform shown to interact with a certain target protein in vitro

or in a heterologous system may not do so in a whole plant context due to spatial and

temporal expression patterns and regulatory mechanisms impacting on the ability of

14-3-3 isoforms to bind to the target protein. Thus, a homologous system is needed

in which 14-3-3 target protein interaction can be tested for, i.e. Arabidopsis 14-3-3

protein interactions will ideally be tested in Arabidopsis or at least in a plant system.

Such a system became available in form of the bimolecular fluorescence

complementation (BiFC) analysis (Bracha-Drori et al., 2004; Walter et al., 2004).

1.8. Bimolecular fluorescence complementation (BiFC):

live cell imaging for protein-protein interaction

Bimolecular fluorescence complementation, developed by Kerppola and

colleagues in 2002, is a very powerful technique to visualise protein interactions in a

native system. It was first used to show protein-protein interactions in living

mammalian cells (Hu et al., 2002; Hu and Kerppola, 2003), but was soon adapted to

fungal and plant systems (Bracha-Drori et al., 2004; Walter et al., 2004; Hoff and Kück,

2005). In plants, BiFC was used to test for the interaction of the Arabidopsis bZIP63

transcription factor with the zinc finger protein LSD1 and to demonstrate 14-3-3 dimer

formation in tobacco (Walter et al., 2004). Now BiFC is one of the most commonly


used techniques for the analysis of protein-protein interactions (reviewed in Kerppola,

2008). The technique is based on the re-association of two non-fluorescent

fragments of the yellow fluorescent protein (YFP). The coding regions of these two

fragments comprising of roughly equal sized amino (N-YFP) and carboxyl terminal

(C-YFP) parts of YFP are translationally fused to the coding regions of two proteins (e.g.

candidate protein 1 and 2), which are to be tested for interaction. The constructs

(candidate protein 1: N-YFP and candidate protein 2: C-YFP) are co-transformed into a

host system, which can be the native system from which the genes encoding the

candidate proteins were isolated. Re-association of a fluorescent YFP complex would

occur when the two candidate proteins interact with each other and the two YFP

fragments are brought closely together. The technique has the great advantage that

it can be used for the fast and relatively simple screening of protein-protein

interactions in transient expression systems. However, the greatest strength of the

system is that it allows for the detailed characterisation of such interactions in the

native biological system. Here, spatial and temporal interaction patterns can be

studied in living organisms, taking into account regulatory factors such as the

availability of light and nutrients or the impact of stresses such as low temperatures

or pathogens.

1.9. Aims of this project

The overall aim of this project was to investigate the functional specificity of

Arabidopsis 14-3-3 proteins with regards to physical protein interactions in vivo using

BiFC analyses. It was first necessary to adapt a BiFC assay system in planta for the

analysis of 14-3-3 interactions. Walter et al. (2004) had previously demonstrated

homodimerisation of a tobacco 14-3-3 isoform, T14-3c, using BiFC analysis in

transiently transformed Arabidopsis protoplasts and Nicotiana benthamiana leaves.


Thus, the same BiFC expression vectors described in Walter et al. (2004) were initially

used to establish the BiFC assay system for the analysis of Arabidopsis 14-3-3

interactions (Chapter 3). For this it was essential to generate a library of Arabidopsis

14-3-3 isoform genes in the BiFC vectors. In parallel, a suitable transient assay

system to study a large number of 14-3-3 dimers had to be established (Chapter 3).

It was proposed that dimerisations among 14-3-3 isoforms and localisation of 14-3-3

proteins may contribute to functional specificity (discussed in Chapter 4, Section 4.1).

Using the transient BiFC assay established in this project, a systematic study was

performed on the dimerisation and subcellular localisation of Arabidopsis 14-3-3

isoforms in living plant cells (Chapter 4). This was extended to the analysis of 14-3-3

dimers in transgenic Arabidopsis plants, allowing the analysis to be performed in a

whole plant context (Chapter 5). Furthermore, a preliminary study was conducted to

test for interactions of 14-3-3 isoforms with a potential 14-3-3 target protein, histone

deacetylase 2C (HD2C) and to establish if this interaction occurs in a 14-3-3

isoform-specific manner (Chapter 4).

During observation of the subcellular localisation of 14-3-3 dimerisations in the

transient BiFC assay, an atypical cytoplasmic distribution pattern – aggregation of BiFC

fluorescence into punctuate bodies – was detected. Attempts to identify possible

organellar destination for the 14-3-3 dimers and possible mechanism for the induction

of 14-3-3 dimer aggregations were conducted (Chapter 6).

In summary, aims of this project were to:

establish an in planta BiFC assay system for the analysis of 14-3-3 interactions

(Chapter 3).

test for dimerisations between Arabidopsis 14-3-3 isoforms and determine

subcellular localisation of the 14-3-3 dimers (Chapter 4).

test for interaction specificity of 14-3-3 isoforms with the potential target HD2C

and subcellular localisation of the 14-3-3-HD2C interaction (Chapter 4).


verify the 14-3-3 dimerisations in a whole plant context using transgenic

Arabidopsis plants (Chapter 5).

identify possible organellar destinations and mechanisms for the induced

aggregation of 14-3-3 dimer into punctuate bodies (Chapter 6).

Chapter 2. Materials and Methods 17

Chapter 2

Materials and Methods


2.1. Materials

2.1.1. General Materials

General chemicals used in this study were of analytical or molecular biology

grade, and were purchased from Amresco (OH, USA), Sigma-Aldrich (MO, USA), Ajax

Finechem (NSW, Australia) and BDH/VWR International Ltd (UK) unless otherwise

stated. Water used in this work for preparation of buffers, solutions and media was

purified using a Milli-Q® Plus Ultra Pure Water System (Millipore, USA) prior to use.

2.1.2. Biological materials

2.1.2.1. Escherichia coli strains used in this study

TOP10: F- mcrA Δ (mrr-hsdRMS-mcrBC) Φ80lacZΔM15 ΔlacX74 recA1 araD139

Δ(araleu)7697 galU galK rpsL (StrR) endA1 nupG (Invitrogen, CA, USA)

DH5α: F- Φ80lacZΔM15 Δ (lacZYA-argF)U169 recA1 endA1 hsdR17(rk-, mk

+) phoA

supE44 thi-1 gyrA96 relA1 λ- (Invitrogen)

2.1.2.2. Agrobacterium strains used in this study

Agrobacterium tumefaciens strain GV3101/pMP90 (Koncz and Schell, 1986)

carrying the pSOUP plasmid was a gift of Dr. Isara Pracharoenwattana and Prof. Steven

Smith, ARC Centre of Excellence in Plant Energy Biology, UWA, Perth, Australia. A.

tumefaciens strain C58C1/pCH32 was kindly given by Dr. Anne Bersoult and Prof. Ian

Small, ARC Centre of Excellence in Plant Energy Biology, UWA, Perth, Australia.

2.1.2.3. Plant materials

Arabidopsis thaliana (Columbia ecotype; Col-0) seeds were purchased from the

Arabidopsis Biological Resource Centre (ABRC).


Nicotiana benthamiana (Nicotiana) seeds were generously provided by Prof.

Mike Jones, Murdoch University, Perth, Australia.

2.1.2.4. Plasmids and primers

Plasmids:

The pUC-SPYNE and pUC-SPYCE plasmids were kindly provided by Profs. Claudia

Oecking, and Klaus Harter, ZMBP, Pflanzenphysiologie, Universität Tübingen,

Tübingen, Germany (Walter et al., 2004).

The pCRII-TOPO® vector was provided with the TOPO TA Cloning® kit

(Invitrogen).

The p19 suppressor of gene silencing plasmid (Voinnet et al., 2003) was a gift of

Prof David Baulcombe at the Sainsbury Laboratory Plant Biosciences Limited, Norwich,

UK.

The pSOUP plasmid and pGreen vectors (Hellens et al., 2000b), pGreen0179 and

pGreen0229, were obtained from John Innes Centre, Norwich, UK

(http://www.pgreen.ac.uk).

The pCAMBIA 1302 plasmid was obtained from CAMBIA, Canberra, ACT,

Australia (http://www.cambia.org).

Constructs encoding four different fluorescent protein-tagged organelle

markers, targeting to the mitochondrion, ER, Golgi or peroxisome, were gifts of Dr.

John Bussell and Prof. Steven Smith (ARC Centre of Excellence in Plant Energy Biology,

UWA, Perth, Australia) and are available through the DNA Stock Centre of the ABRC

(http://www. arabidopsis.org). These organelle markers, comprising of a

multicoloured set, were generated by Nelson and co-workers (Nelson et al., 2007).

The mitochondrial, ER and Golgi markers, tagged with mCherry fluorescent protein, a

http://www.cambia.org/


derivative of the monomeric red fluorescent protein, mRFP1, (Shaner et al., 2004),

were used in this study. The mitochondrial marker was created by incorporating the

transit peptide sequence of the yeast mitochondrial cytochrome c oxidase IV (ScCOX4;

Köhler et al., 1997) upstream of the mCherry sequence. The ER targeting marker

was made by adding the signal peptide of the Arabidopsis cell wall-associated kinase

2 (AtWAK2; He et al., 1999) at the N-terminus and the ER retention signal

His-Asp-Glu-Leu (HDEL) at the C-terminus of the mCherry sequence (Gomord et al.,

1997). The Golgi targeting marker was generated by integrating the first 49 amino

acids of soybean α-1, 2-mannosidase I (GmMan1; Saint-Jore-Dupas et al., 2006) at the

N-terminus of the mCherry marker. All of these marker genes are under the control

of a double 35S promoter and carried by the pBIN-based binary plasmid (Nelson et al.,

2007).

The peroxisome marker used in this study is genetically engineered by addition

of the peroxisomal targeting signal 1 (PTS1) sequence (IHHPRELSRL) of pumpkin

malate synthase (MLS) at the C-terminus of the mRFP (Pracharoenwattana et al.,

2005). The peroxisome marker construct is driven by a double 35S promoter

(Pracharoenwattana et al., 2005).

Primers:

Primers used in this study are summarised in the following tables. The primers

used for the amplification of the open reading frame (ORF) of twelve 14-3-3 isoform

genes are listed in Table 2-1. The primers used for general purposes are listed in

Table 2-2.

2.2. Nucleic acid methods

All molecular cloning manipulations and procedures were done according to the

standard protocols in Sambrook et al. (1989) unless otherwise stated.


Table 2-1. Primers for amplification of full-length coding regions of 14-3-3s

or the coding regions of N-terminally truncated (N-Δ) 14-3-3s.

14-3-3 Forward primer (for full-length) Reverse primer* Forward primer (for N-Δ)

chi 5’-CAACAATGGCGACACCAGGAG 5’-tcgcctcgagGGATTGTTGCTCGTCAGC 5’-ATGGAGAAAGTCGCGAAAG

epsilon 5’-CTATGGAGAATGAGAGGGAAAAGC 5’-tcgcctcgagGTTCTCATCTTGAGGCTCATC 5’-ATGAAGAAAGTTGCTCAGCTTG

iota 5’-ACAAAATGTCATCATCAGGATCCG 5’-tcgcctcgagGTTCTCAGTGGCATCGG 5’-ATGAAGAAAGTGGCGAGG

kappa 5’-CTCTAATGGCGACGACCTTAAGC 5’-tcgcctcgagGGCCTCATCCATCTGCTC 5’-ATGGAACAGCTCGTAAGTG

lambda 5’-CTCTAATGGCGGCGACATTAGG 5’-tcgcctcgagGGCCTCGTCCATCTGCTC 5’-ATGGAACAGCTCGTTACAG

mu 5’-CAGTCATGGGTTCTGGAAAAGAG 5’-tcgcctcgagCTCTGCATCGTCTCCAC 5’-ATGAAAAGTGTTGCGAAATTAAATG

nu 5’-ATAAGATGTCGTCTTCTCGGGAAG 5’-tcgcctcgagCTGCCCTGTCTCAGCTG 5’-ATGGAGAAAGTTGCAAAGAC

omega 5’-CAACAATGGCGTCTGGGCGTG 5’-tcgcctcgagCTGCTGTTCCTCGGTCG 5’-ATGGAGAAAGTCTCCGCC

omicron 5’-CTATGGAGAACGAGAGAGCGAAGC 5’-tcgcctcgagCTTACCTCCTTCCTCTAGATC 5’-ATGAAGAAAGTTGCAGCTCTTG

phi 5’-TAATGGCGGCACCACCAGCATC 5’-tcgcctcgagGATCTCCTTCTGTTCTTCAGC 5’-ATGGAAAAAGTCGCTGAAGC

psi 5’-CGAAGATGTCGACAAGGGAAGAG 5’-cgccctcgagCTCGGCACCATCGGG 5’-ATGGAGAAAGTTGCGAAAACTG

upsilon 5’-CAAAGATGTCTTCTGATTCGTCCCG 5’-tcgcctcgagCTGCGAAGGTGGTGG 5’-ATGGAGAAAGTTGCAAAGACC

* Artificially designed sequence adaptors are in lower case. The first ATG for the 14-3-3 ORFs is

shown in bold. The XhoI restriction site sequence (ctcgag), which replaced the nucleotides

encoding the stop codons of 14-3-3 isoforms, is underlined.

Table 2-2. Primers used for general purposes.

Primer name Primer sequences Description

35S-Seq (forward)

5’-GTAAAGACTGGCGAACAG For sequence determination of 14-3-3 ORFs in the pGreen-based BiFC vectors and for PCR genotyping

NYFPsp1 (reverse)

5’-ATGAACTTCAGGGTCAGC as above

CYFPsp1 (reverse)

5’-AGCTCAGGTAGTGGTTGTC as above

2.2.1. Plasmid DNA isolation

2.2.1.1. Plasmid DNA preparation

All of the following procedures were performed at room temperature. A single

transformant colony was picked from selective Luria-Bertani Broth [LB; Sambrook et

al. (1989)] plates and grown overnight at 37°C in liquid selective LB with shaking at


225 rpm. Cells were collected by centrifugation (12000 x g; Eppendorf centrifuge

5415D, Hamburg, Germany) and re-suspended in Miniprep Solution I (Appendix I).

Miniprep Solution II (Appendix I) was added to lyse the cells, and the suspension was

then neutralised by adding Miniprep Solution III (Appendix I). The cell debris was

collected at 12000 x g for 5-10 min and the supernatant was transferred to a new

microfuge tube. Ice-cold 100% ethanol was added into the supernatant to

precipitate the plasmid DNA. After a centrifugation at 12000 x g for 2 min, the

plasmid DNA was washed in 70% ethanol, air-dried and re-suspended in sterile Milli-Q

H2O (MQ H2O).

2.2.1.2. Isolation of plasmid DNA using a commercial kit

Highly purified plasmid DNA was used for automated DNA sequence

determination (Section 2.2.7), transformation of bacteria (Section 2.3.2 and 2.3.4) and

polyethylene glycol (PEG)-mediated transfection of Arabidopsis mesophyll protoplasts

(Section 2.4.3). High purity plasmid DNA was isolated using a commercial miniprep

kit, the Wizard® Plus SV Minipreps DNA purification System (Promega Co., WI, USA)

according to the manufacturer’s instructions.

2.2.2. Restriction enzyme digestion

The restriction enzymes used in this study were purchased from either TaKaRa

Bio Inc. (Shiga, Japan), New England Biolabs Inc. (MA, USA) or Promega. A typical

restriction digestion procedure was done by using 0.5-2 μg of plasmid DNA digested

with 2-10 units of restriction enzyme in 1 x supplied enzyme buffer to a final volume

of 20 μl. Sometimes bovine serum albumin (BSA) was added, according to

manufacturer’s instruction. RNase A (0.5-1 μg; Invitrogen) was added to digest RNA

molecules in the plasmid DNA preparation (section 2.2.1.1). Temperature and


incubation time for restriction digests were used as recommended by the suppliers.

The digested plasmids were analysed by agarose gel electrophoresis (section 2.2.3).

2.2.3. Agarose gel electrophoresis

In general, 0.6%-2% agarose TAE (Appendix I; Sambrook et al., 1989) gels

containing 0.5 μg ml-1 ethidium bromide (EtBr) were used to separate linear DNA

fragments, depending on their molecular sizes. DNA samples were mixed with

one-fifth (1/5) volume of Orange G loading dye (Appendix I) before loading into gel

wells. Samples were separated in 1x TAE buffer in an electrophoresis apparatus

(SUB-CELL® GT or MINI-SUB CELL® GT, Bio-Rad Laboratories Pty., Ltd., NSW, Australia)

at a constant voltage between 50 and 100 V, depending on the percent agarose, until

the dye migrated to the bottom of the gel. Images were taken using a Gel Doc™ XR

and ChemiDoc™ XRS Gel Documentation System and Quantity One software (Bio-Rad),

and then were processed using Adobe Photoshop® 5.0 software (Adobe System Inc.,

CA, USA).

2.2.4. Ligation of DNA fragments into vectors

For subcloning purposes, DNA fragments (inserts and vectors) were purified

from agarose gels using the Wizard® SV Gel and PCR Clean-up System (Promega). In

general, the purified inserts were mixed with the destination vectors in a molar ratio

of 3:1 for ligation, and with 0.1-1 unit of T4 DNA ligase (Promega) and 1 x ligase buffer

in a total volume of 10 μl. The ligation reaction was incubated at 14°C overnight.

The recombinant plasmids were then transformed into chemically competent

Escherichia coli (E. coli) cells (Section 2.3.2).


2.2.5. Polymerase Chain Reaction (PCR)

A typical 25 μl PCR mixture for an analytical PCR included: 1 x NH4 Reaction

Buffer (Bioline), 2 mM MgCl2, 1 mM dNTP mix (250 μM each), 0.25 μM forward and

reverse primers, and 0.625 unit of Taq polymerase (BIOTAQ®, Bioline). The amount

of required template depended on the source of DNA sample. Generally, picogram

levels of plasmid DNA or microgram levels of genomic DNA were required as a PCR

template.

A typical thermocycling program generally used for amplification of an 1 kb DNA

fragment was as follows: 96°C for initial denaturation for 5 min, 30-35 cycles of 96°C

for 30 s, 55°C (for primers annealing) for 30 s and 72°C (for elongation) for 1 min,

followed by a final elongation step at 72°C for 6 min. PCR was done using a

programmable thermal cycler (PTC-100, MJ Research Inc., MA, USA). PCR products

were analysed by agarose gel electrophoresis (Section 2.2.3).

In order to minimise the possibility of PCR errors, a high-fidelity Taq polymerase

(Accuzyme®, Bioline) was used to amplify coding sequences of Arabidopsis 14-3-3

isoforms. The DNA templates used for the amplification were from either an

Arabidopsis cDNA library (Minet et al., 1992), a mixture of first strand cDNA

generated from Arabidopsis leaves and roots (Dr. T. Martin laboratory), or 14-3-3

cDNA clones purchased from the ABRC. The primers used in the amplification

reactions are listed in Table 2-1. The thermocycling program used for the

high-fidelity Taq polymerase, Accuzyme®, was similar to a typical PCR, except a longer

elongation time was used (1.5-2 min for 1 kb).

2.2.6. TOPO TA Cloning®

The PCR products encoding DNA fragments of interest were cloned into the

pCRII-TOPO® vectors using the TOPO TA Cloning® kit (Invitrogen) as per the

manufacturer’s instructions.


2.2.7. Sample preparation for automated DNA sequence determination

The BigDye® Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems) was

used for automated DNA sequence determination. Purified plasmid DNA or PCR

fragments were used as template for the sequencing reactions. A sequencing

reaction was prepared by mixing 2 μl of BigDye® Terminator reaction mix, 2 μl of 2.5 x

sequencing buffer (Applied Biosystems), 100-250 ng of double-stranded DNA and 1.6

pmol of sequencing primer in a final volume of 10 μl. Cycle sequencing was done

with a thermal cycler (PTC-100, MJ Research) using the following program: an initial

denaturation at 96°C for 1 min, followed by 30 cycles of 96°C for 10 s, 50°C for 5 s and

60°C for 4 min. Samples were added to 1 μl of 3 M sodium acetate, pH 4.6-4.7 and

25 μl of 95% ethanol for DNA precipitation. After 15 min, the sample was collected

at 16000 x g for 20 min, washed gently with 70% ethanol and then collected again at

16000 x g for 5 min. After drying, the samples were submitted to the Lotterywest

State Biomedical Facility Genomics (LSBFG), Perth, Australia

(http://lsbfg.meddent.uwa.edu.au/) for sequence determination.

2.3. Bacterial methods

2.3.1. Preparation of competent E. coli cells

The rubidium chloride method for preparation of competent cells was used as

described by Hanahan (1985). An E. coli strain (section 2.1.2.1) was streaked on a LB

plate and grown overnight at 37°C. A single colony from the plate was used to

inoculate 3 ml LB liquid medium and the bacteria were grown overnight at 37°C with

shaking at 225 rpm. An aliquot of the starter culture was used to inoculate a 100 x

volume of LB medium containing 20 mM MgSO4. Cells were grown at 37°C with

shaking at 225 rpm until the optical density at 600 nm (OD600) of the culture reached

0.4-0.6. The cells were collected by centrifugation at 4500 x g for 5 min at 4°C.


Cells were kept at 4°C or on ice for the rest of the procedure and all of the following

preparation steps were performed in a cold room. The cell pellet was gently

re-suspended in a 0.4 volume of ice-cold TFB 1 (Appendix I). Cells were incubated

on ice for 5 min and subsequently collected by centrifugation at 4500 x g for 5 min at

4°C. The supernatant was decanted and the cell pellet was re-suspended very gently

in 1/25 volume of the subculture volume using ice-cold TFB 2 (Appendix I). The cell

suspension was incubated on ice for 15-60 min, and then divided into 100 or 200 μl

aliquots, snap-frozen in liquid nitrogen and then stored at -80°C.

2.3.2. Transformation of E. coli competent cells

Competent cells were thawed on ice. An aliquot (minimum 10 μl) of

competent cells was mixed gently with 1 μl of plasmid DNA (approximately 10 ng of a

ligation reaction). The suspension was incubated on ice for 30 min. The cells were

heat shocked in a 42°C water bath for 45-60 s, followed by incubation on ice for 2 min.

LB medium (150 μl) was added to the cell suspension and the cells were incubated at

37°C for 1 h with shaking (225 rpm). A 20 μl aliquot of the transformed cells was

spread onto a selective LB plate with appropriate antibiotic(s) or selection of

transformants. The remainder of the transformation mix was spread on a second

selective LB plate. The cells were incubated overnight at 37°C.

For selection of E. coli cells carrying pCRII-TOPO-based recombinant vectors,

ampicillin (100 μg ml-1) or kanamycin (50 μg ml-1) was used. For E. coli cells carrying

the pCAMBIA 1302 vector or pGreen-based vectors, kanamycin (50 μg ml-1) was used.

2.3.3. Preparation of competent Agrobacterium cells

The A. tumefaciens strain GV3101/pMP90 containing the pSOUP plasmid was

streaked and grown on a LB plate with rifampicin (50 μg ml-1), gentamicin (5 μg ml-1)


and tetracycline (2 μg ml-1) at 28-30°C for 2-3 days. A single colony was used to

inoculate 3 ml LB medium with rifampicin (50 μg ml-1) and gentamicin (5 μg ml-1) and

cells were grown at 28-30°C with 225-250 rpm shaking overnight. An aliquot (1 ml)

of the overnight culture was used to inoculate 100 ml LB medium containing

rifampicin and gentamicin and the cells were grown at 28-30°C with 225-250 rpm

shaking until the culture reached an OD680 of 0.5-1.0. The culture was chilled on ice

for 10 min, then transferred to pre-chilled tubes and cells were collected at 3000 x g

for 10 min at 4°C. The cell pellet was rinsed with 1 ml of ice-cold 20 mM CaCl2 and

collected again by centrifugation to remove the residual antibiotics. The cell pellet

was re-suspended in 2 ml ice-cold 20 mM CaCl2 (1/50 volume of culture) and 0.2 ml

aliquots were snap-frozen in liquid nitrogen and stored at -80°C.

2.3.4. Transformation of Agrobacterium cells

The freeze-thaw method described here for transformation of Agrobacterium

cells was an adaptation from the protocol of Höfgen and Willmitzer (1988).

Competent Agrobacterium cells were thawed on ice. A 20 μl aliquot of the cells was

mixed with 1-2 μl of purified plasmid DNA, incubated on ice for 5 min, frozen in liquid

nitrogen for 5 min and then thawed in a 37°C water bath for 5 min. LB medium (100

μl) was added and the cell suspension was mixed. The suspension was incubated at

28-30°C with shaking at 250 rpm for 2 h. The cells were spread on LB plates

containing appropriate antibiotics (Table 2-3) and grown at 28-30°C for 2-3 days.

Positive clones were confirmed using colony PCR analysis (Section 2.3.5).

2.3.5. Colony PCR

Transformed Agrobacterium cells were confirmed by PCR. A single

transformant colony was re-suspended in 10 μl sterile MQ H2O. A 5 μl aliquot of the


cell suspension was used as the template for a general PCR (Section 2.2.5) with

specific primers for the target sequence in the recombinant plasmids. The remaining

5 μl of the suspension was used to make a glycerol stock (Sambrook et al., 1989) for

subsequent use.

Table 2-3. Antibiotics used for selection of Agrobacterium cells.

Agrobacterium

strain

for chromosomal

selection

for Ti-plasmid

selection

for binary vector

selection

for helper plasmid

selection

GV3101 rifampicin

(50-100 μg ml-1)

pMP90

(gentamycin 5-10 μg ml-1)

pGreens or

pCAMBIA 1302

(kanamycin 50 μg ml-1)

pSOUP

(tetracycline 2 μg ml-1)

C58C1 rifampicin

(50-100 μg ml-1)

pCH32

(tetracycline 2 μg ml-1)

p19 vector

(kanamycin 50 μg ml-1) -

2.4. Plant methods

2.4.1. Plant growth

In general, A. thaliana and N. benthamiana plants were grown on compost mix,

which was prepared using 70% volume of Shamrock® composts (Seed & Modular

propagating medium, Scotts, UK) with 30% of fine vermiculite from a local supplier

(Perlite & Vermiculite Factory, WA, Australia). Plants were grown in a growth cabinet

at 22 ± 1°C under a long day photoperiod (16 h of approximately 100 μE m-2 s-1 light

and 8 h dark) unless otherwise stated. Water was provided from the bottom of pots.

For germination, Arabidopsis seeds were stratified at 4°C in the dark for 2 days before

transfer into the growth cabinets; while Nicotiana seeds did not need stratification for

germination.


2.4.2. Isolation of Arabidopsis mesophyll protoplasts

The following procedures to isolate mesophyll protoplasts from Arabidopsis

leaves were combinations of protocols from Huang and Chen (1988) and Yoo et al.

(2007), with minor modifications to adapt to facilities available in the laboratory.

Mesophyll protoplasts were prepared from rosette leaves of 4-5 week-old

soil-grown Arabidopsis plants. Approximately 0.5 g of leaves were detached and

briefly surface sterilised in 70% ethanol for 1 min, followed by three washes with

sterile MQ H2O. The leaves were sliced into 1-2 mm wide strips in 10 ml of

protoplasting solution [1% cellulase (onozuka R-10; Duchefa Biochemie, Netherlands),

0.2% macerozyme (Duchefa Biochemie), 0.4 M sucrose, 20 mM KCl, 10 mM CaCl2, 20

mM 2-(N-morpholino) ethanesulfonic acid (MES)-KOH, pH 5.7, 0.45 μm filtered]

within a 9 cm Petri dish, and then vacuum-infiltrated for 25 min to allow the

protoplasting solution to permeate the intercellular airspaces. Subsequently, the

leaf strips were incubated at room temperature in the dark for 3-16 h to give the

enzymes sufficient time to digest cell walls. The release of protoplasts gradually

turned the yellowish, clear protoplasting solution into greenish over time. The

greenness of the solution and the digested leaf strips were used as primary indicators

for the release of protoplasts. The protoplast suspension was passed through a

sieve (approximate 1 mm pore) to remove the large pieces of undigested debris.

The crude protoplast suspension was carefully transferred to a 15 ml Falcon tube

using a wide-bore pipet, and the cell debris was pelleted by centrifugation at 100 x g

for 5 min using a swing bucket centrifuge (Eppendorf centrifuge 5804). The floating

protoplasts (as a thin, green layer at the top of supernatant) were transferred

(approximately 1 ml) to a 15 ml tube. The volume was brought to 10 ml with W5

solution (154 mM NaCl, 125 mM CaCl2, 5 mM KCl, 2 mM MES-KOH, pH 5.7) and the

suspension then mixed gently by inversion. The protoplasts were collected as


described above. The supernatant was removed using a wide-bore pipet and the

protoplast pellet was re-suspended 5 ml of fresh W5 solution.

2.4.3. Polyethylene glycol-mediated transfection of A. thaliana mesophyll

protoplasts

The number of protoplasts was estimated by diluting the suspension at least

100 x in W5 solution. Three aliquots of 10 μl of diluted protoplasts were placed on a

glass slide and the number of protoplasts was counted using an inverted microscope

(Olympus IX71). To assess protoplast viability, a 50 μl aliquot of protoplast

suspension was stained with 400 μg ml-1 methylene blue (Huang et al., 1986).

Damaged protoplasts and contaminating cell debris were stained blue, whereas viable

protoplasts excluded the dye and remained green. More than 90% of the

protoplasts were viable in a typical preparation.

This PEG-mediated transfection of protoplasts followed the protocol described

by Yoo et al. (2007). The protoplast suspension (Section 2.4.2) was incubated on ice

for at least 30 min. The chilled protoplasts were collected at 100 x g for 5 min and

gently re-suspended to a density of 2 x 105 cells ml-1 in MaMg solution (0.4 M

mannitol, 15 mM MgCl2, 5 mM MES-KOH, pH 5.7). A 100 μl aliquot of protoplast

suspension (approximate 2 x 104 protoplasts) was mixed with 10 μl of purified

plasmid DNA (10-20 μg plasmid DNA) by gentle swirling using a pipet tip.

PEG-calcium solution [110 μl; 40% (w/v) PEG 3350 (Sigma), 0.2 M mannitol, 100 mM

CaCl2; Appendix I] was added to the protoplast-DNA mixture and then the entire

suspension was mixed to homogeneity by gently swirling. The suspension was

incubated at room temperature for 5-30 min, followed by the addition of 0.44 ml of

W5 solution and gentle mixing. The protoplasts were collected again at 100 x g for 5

min and the pellet was diluted gradually with W5 solution to a final volume of 1 ml.


The transfected protoplasts were incubated in a growth cabinet at 22°C in the dark.

After 16-24 h incubation, the protoplasts were analysed for transgene expression.

For the BiFC analysis of 14-3-3 dimerisation, equal amounts of a 14-3-3-YN and a

14-3-3-YC plasmid (Section 3.2.4 and Table 3-2) were combined for the transfection of

Arabidopsis mesophyll protoplasts following the procedures described above. In

parallel using a second protoplast sample, a green fluorescent protein (GFP)

gene-containing plasmid, pCAMBIA 1302, was used as a positive control to evaluate

the efficiency of the transfection method. Fluorescence was examined 24-48 h after

transfection using a conventional fluorescence microscope (Section 2.5.1).

2.4.4. Agroinfiltration of Arabidopsis and Nicotiana leaves

The procedures for agroinfiltration of Arabidopsis leaves were as previously

described in Bracha-Drori et al. (2004). The agroinfiltration of Nicotiana leaves was

based on several previous studies (Voinnet et al., 2003; Walter et al., 2004; Schütze et

al., 2009) with minor modifications. Briefly, a colony of the Agrobacterium strain

harbouring a plant expression vector was grown in 3 ml of LB medium with

appropriate antibiotics (Table 2-3). Cells were grown overnight at 28-30°C with

shaking (225 rpm). The cells of the overnight culture were collected at 16000 x g for

30 s and re-suspended in 1 ml of infiltration buffer without acetosyringone (10 mM

MgCl2, 10 mM MES-KOH, pH 5.6). Then the cell suspension was diluted to an OD600

0.7-0.8 with the infiltration buffer containing 200 μM acetosyringone and incubated

at room temperature for 1 h. Working suspensions for the infiltration were

prepared by mixing desired Agrobacterium clones for the transient assay.

For the BiFC assay, the working suspensions were prepared by mixing

appropriate clones containing the BiFC constructs and the p19 plasmid at a 1:1:1 ratio.

For a GFP positive control, the pCAMBIA 1302 (35S-GFP)-containing Agrobacterium


clone was combined with the p19 clone for leaf infiltration. In addition, the working

suspensions for the co-localisation study of 14-3-3 dimeric complexes and organelle

markers (Section 2.1.2.4) were prepared by mixing appropriate clones containing the

BiFC constructs, the p19 plasmid and an organelle marker construct at a 1:1:1:0.1

ratio (Section 4.2.5).

Infiltration mixtures were injected into the abaxial air space of leaves of 6-8

week-old Nicotiana plants using a 1 ml syringe (without a needle) until the entire leaf

air space was filled up with the Agrobacterium suspension. The injected leaves were

marked with a pen. Lower epidermal layers of infiltrated leaves were examined for

fluorescence three days after infiltration using a fluorescence microscope or a

confocal microscope (Section 2.5.1 and 2.5.2).

2.4.5. Transformation of Arabidopsis using floral dipping

The floral dip method for Arabidopsis transformation was an adaptation of

protocols of Clough and Bent (1998) and Logemann et al. (2006). Growth of

Arabidopsis plants was as previously described (Section 2.4.1). Primary bolts of

healthy, flowering Arabidopsis plants were clipped to trigger the growth of more

flowering branches. The floral dip transformation was performed when the new

flowering branches grew to approximately 10 cm in height and contained some open

flowers and few green siliques (approximately 4-6 days after clipping).

A colony of transformed Agrobacterium cells was completely re-suspended in

20 μl sterile MQ H2O. The bacterial suspension was evenly spread on the entire

surface of a LB plate containing appropriate antibiotics (Table 2-3). The bacteria

were grown at 28-30°C for 2-3 days until a confluent bacterial lawn was obtained. To

collect the densely grown bacteria, approximate 4 ml of 5% sucrose were added to

the plate, and the bacteria were scraped off from the agar and suspended in the


sucrose solution. The bacterial suspension was then transferred to microfuge tubes.

The cells were collected at 4000 x g for 10 min at room temperature and the

supernatant was decanted. The pellets were re-suspended in 5% sucrose containing

0.01% silwet L-77 to reach an OD600 of 0.8 for the floral dipping. The suspension was

poured into a disposable plastic bag and the inflorescences of plants (approximately

20 plants per pot) were dipped into the solution for 30 s with gentle agitation. The

inflorescences were covered loosely by placing a plastic bag over the pot for 16 to 24

h to maintain high humidity in the micro-environment surrounding the dipped flowers.

A second dip was performed seven days after the first dip. The plants were then

grown under the conditions described in Section 2.4.1 until seeds matured. Mature

seeds were collected and screened for transformation events using appropriate

antibiotics or herbicides selection methods (Section 2.4.7).

2.4.6. Surface sterilisation of Arabidopsis seeds

Approximately 20 mg of seeds were mixed with 1 ml of seed sterilisation

solution (Appendix I) in a microfuge tube and shaken vigorously for 5 min. Seeds

were then washed two times with 1 ml of 95% ethanol. After each wash, ethanol

was removed using a micro pipette. The seeds were left in a laminar flow hood for a

few hours to evaporate residual ethanol.

To surface sterilise a small amount of seeds (20-50 seeds), seeds were

dampened with 70% ethanol on a sterile filter paper in a laminar flow hood, followed

by rinsing the seeds with 95% ethanol. When the ethanol completely evaporated,

the dry, sterilised seeds were sprinkled on the surface of a 0.5 x Murashige and Skoog

(MS) medium plate (Appendix I).


2.4.7. Screening Arabidopsis transformants

2.4.7.1. Screening transformants on plates

Arabidopsis seeds harvested after floral-dip transformation (T1 seeds) were

screened for transformants using antibiotics or herbicide, depending on the plant

selection marker gene carried in the binary vector. For the pGreen0179-based

14-3-3 BiFC constructs, hygromycin B was used; while Basta® (glufosinate ammonium,

a herbicide) was used for the pGreen0229-based constructs. The selection procedures

for transformants were done according to Harrison et al. (2006). Sterilised T1 seeds

(Section 2.4.6) were sprinkled on 0.5 x MS medium plates containing 30-50 μM Basta

or 10-15 μg ml-1 hygromycin B, and then stratified at 4°C in the dark for 2 days.

After stratification, the plates were transferred to growth cabinets and incubated for

4-6 h at 22°C in continuous white light to stimulate seed germination. The plates

were then wrapped with aluminium foil to protect the seeds from light, and

incubated at 22°C for 2 days. After the foil was removed, germinated seedlings were

grown at 22°C in continuous white light for 24-48 h or in a 16 h day:8 h night

photoperiod for the selection of phenotypes. Transformants were screened based

on the following phenotypes: Basta-resistant seedlings would have long hypocotyls

and green cotyledons, non-resistant seedlings would have long hypocotyls and pale

cotyledons. In contrast, seedlings resistant to hygromycin B would have long

hypocotyls, green cotyledons and elongated roots, whereas non-resistant seedlings

would have short hypocotyls and short roots, but had also green cotyledons.

Candidate transformants were transferred to pots containing compost:vermiculite

mix (Section 2.4.1) and transgenes were confirmed by PCR genotyping (Section 2.4.9)

once leaves had developed.

2.4.7.2. Screening for transformants using Basta selection on soil-grown

plantlets


Basta resistant transgenic Arabidopsis plants can also be selected for when

grown in soil (Weigel and Glazebrook, 2002). To do this, T1 seeds were sown on

pre-dampened compost mix (Section 2.4.1) and then stratified at 4 oC in the dark for 2

days. Pots were removed from 4 oC conditions and transferred to growth cabinets

for germination and growth of seedlings. Once green cotyledons of seedlings were

fully expanded, seedlings were sprayed with Basta solution (200 μM in water) 2-3

times over a two-day period. Cotyledons of transformants stayed green and such

seedlings continued to grow. Cotyledons of untransformed plants and wild type

controls turned yellow and wilted with no further growth. Potential transformants

were transferred individually to pots with new compost mix and grown until they

developed several leaves. Leaves were used to confirm positive transformation

events by PCR genotyping (Section 2.4.9).

2.4.8. Plant genomic DNA isolation

2.4.8.1. Genomic DNA isolation using the cetyltrimethyl ammonium

bromide (CTAB) method

The CTAB method to isolate plant genomic DNA was performed according to

Gawel and Jarret (1991). Approximately 10 mg of leaf tissues were ground at room

temperature with 500 μl CTAB extraction buffer (Appendix I). The CTAB extraction

buffer was added in two aliquots: initially 50 μl were added to initiate the grinding

process. The remaining buffer was added when most of the tissue was separated

into smaller pieces. The sample was incubated at 65°C for 2 h with frequent mixing

and then cooled to room temperature. Following the addition of 4 μg of RNase A

(Invitrogen), the sample was incubated at 37°C for 1 h. The sample was extracted

twice with an equal volume of chloroform:isoamyl alcohol (24:1, v/v). The final

aqueous phase was added to two-thirds volume of 100% isopropanol and mixed


gently to precipitate the DNA at -20°C overnight. The DNA was collected at 16000 x

g at room temperature for 15 min and washed twice with two-thirds volume of 70%

ethanol. The DNA pellet was air-dried at room temperature for 5-20 min and then

re-suspended in 50 μl of sterile MQ H2O. A 5 μl aliquot was used to check the

quality of DNA by agarose gel electrophoresis (Section 2.2.3).

2.4.8.2. Genomic DNA isolation using the method of Wang et al. (1993)

Genomic DNA from plants prepared using the CTAB method (section 2.4.8.1)

was of a high quality and well suited for PCR characterisation. Alternatively, a faster

protocol described by Wang et al. (1993) was used to obtain genomic plant DNA for

PCR analysis if the quality of the DNA was not required to be so high, and it is briefly

described as below. One to two pieces of leaves (a few milligrams) were ground in

0.5 N NaOH (5 μl mg-1 leaf tissue) using a microfuge tube pestle until no large pieces

of tissues remained. A 10 μl aliquot of the extract was mixed with 490 μl of 100 mM

tris (hydroxymethyl) aminomethane (Tris)-Cl pH 8.0. An aliquot (5 μl) of the diluted

extract was used directly as a template for an analytical PCR (Section 2.2.5).

2.4.9. PCR genotyping of transgenic plants

The existence of a transgene in a putative transgenic plant was determined by

using microgram amounts of plant genomic DNA (Section 2.4.8) as the template in

combination with specific primers for the gene of interest in a typical PCR (Section

2.2.5), followed by agarose gel electrophoresis (Section 2.2.3).

2.4.10. Biochemical treatments

2.4.10.1. Hydrogen peroxide treatment

Protoplasts were isolated from rosette leaves of transgenic Arabidopsis showing

GFP or BiFC fluorescence (Section 2.4.2). Aliquots of the protoplast suspension


(approximately 2 x 103 protoplasts each) were individually treated with 0, 0.2 and 1

mM Hydrogen peroxide (H2O2) at room temperature and in the dark. Fluorescence

in the protoplasts was examined at 12 and 24 h post-treatment using conventional

fluorescence microscopy.

2.4.10.2. Methyl jasmonate treatment

Aliquots from the same batch of protoplast suspension used in the H2O2

treatment (Section 2.4.10.1) were used for the methyl jasmonate (MeJA) treatment.

Protoplasts were treated with 0.1% ethanol (solvent of MeJA as a mock-treated

control; 0 μM of MeJA) 5, or 50 μM MeJA at room temperature in the dark.

Fluorescence in the protoplasts was examined at 12 and 24 h post-treatment using

conventional fluorescence microscopy.

For MeJA treatment of leaves, detached rosette leaves from 5-6 week-old

transgenic Arabidopsis plants expressing GFP or 14-3-3-BiFC constructs were

sub-immersed in 0.1% ethanol (mock-treated control) or in 50 μM MeJA solution for

38 h. Effects of the MeJA treatment on fluorescence distribution in epidermal cells

were examined by conventional fluorescence microscopy.

For MeJA treatment of seedlings, ten day-old transgenic seedlings with GFP or

BiFC fluorescence were transferred from hygromycin selective plates (1/6 x MS, 1%

sucrose, 15 μg ml-1 hygromycin B) to 0 μM or 50 μM MeJA containing plates (1/6 x MS,

1% sucrose) for 4 days. Absolute root growth during this period was measured.

Epifluorescence images were taken of newly grown tissues (first pair of leaves or

primary root tips) at the end of treatment using conventional fluorescence

microscopy.


2.5. Microscopy and imaging

2.5.1. Fluorescence microscopy

Leaf discs were removed from the Agrobacterium injected area of leaves

(Section 2.4.4) and mounted on glass slides in water for fluorescence microscopy.

For higher resolution of the intracellular distribution of expressed fluorescent

proteins, peels of the lower leaf epidermis were prepared. Both types of

preparations were examined using an Olympus IX71 fluorescence microscope

equipped with a DP70 CCD camera (Olympus). Filter sets (Olympus) used for the

observation of fluorescence in this study are summarised in Table 2-4.

Table 2-4. Filter sets used in fluorescence microscopy in this study.

Information regarding the characteristics of the listed filter sets is based on the Olympus

manual for the IX71 fluorescence microscope.

Mirror Unit Excitation filter

Dichroic mirror

Barrier filter Fluorochromes

U-MWU2 300-385 nm 400 nm 420 nm DAPI

U-MGFPHQ 460-480 nm 485 nm 495-540 nm GFP (and BiFC fluorescence in this

study)

U-MWIB3 460-495 nm 505 nm 510 nm FITC (showing concurrently BiFC

and chlorophyll autofluorescence)

U-MRFPHQ 535-555 nm 565 nm 570-625 nm RFP (for detection of expression of

organelle markers in this study)

Images were captured using the DPController software (Olympus), and then

processed using Adobe Photoshop 5.0® software. In general, the contrast and

dark/light levels of images were manipulated by Photoshop for optimal image

presentation.

2.5.2. Confocal microscopy

Leaf discs from the infiltrated area of Nicotiana leaves or the leaves of transgenic

Arabidopsis expressing 14-3-3 BiFC constructs were placed in water on glass slides and


covered with coverslips for Confocal Laser Scanning Microscopy (CLSM) using a TCS SP2

AOBS Confocal Microscope (Leica). Mesophyll protoplasts were analysed by CLSM

directly in W5 solution on glass slides and covered with coverslips (with little paper

strips as spacers in between coverslips and glass slides to create space for protoplasts).

Confocal images were collected using the Leica Conical Software with appropriate

settings for excitation and emission channels as follows:

GFP: 488 nm for excitation; 510-540 nm for emission.

YFP: 514 nm for excitation; 520-550 nm for emission.

mCherry: 561 nm for excitation; 580-610 nm emission.

Chlorophyll autofluorescence: 514 nm for excitation; 660-700 nm for emission.

Confocal microscopy enables the detection of fluorescence in optical section

scans through a tissue or cell. Projection images of serial optical sections were

compiled and processed using the default setting of the Leica Conical Software. Some

of the confocal images (single optical sections or projection images) were further

processed for contrast or dark/light level manipulations using Adobe Photoshop 5.0®

software.

2.5.3. Image analysis

2.5.3.1. Quantification of fluorescence intensity using ImageJ

The analysis of the subcellular localisation patterns of 14-3-3 dimers in transiently

transformed Nicotiana leaf epidermal cells (Chapter 4, Section 4.2.3.3) was based on

BiFC fluorescence intensity in the nucleus and cytoplasm. These intensities were

determined from obtained micrographs using ImageJ (http://rsbweb.nih.gov/ij/). In

detail, five circular, 10 pixel areas were selected within the nucleus and also within the

cytoplasm. Great care was taken to place these selections within areas showing

representative fluorescence of the individual dimers and cells. The mean grey values


were obtained for the nuclear and cytoplasmic areas using ImageJ. Values obtained in

a similar way for the non-fluorescent background were zero in each case. Thus, the

mean values for nuclear and cytoplasmic signal were used directly to calculate the

nuclear to cytosolic signal ratio for each cell. The ratios were then used to generate

scatter plot diagrams for further analysis of subcellular localisation patterns of 14-3-3

dimers (Fig. 4-3 to 4-5).

2.5.3.2. Image deconvolution

Epifluorescence images were deconvolved using Huygens Essential software

(version 3.4, Scientific Volume Imaging B.V., The Netherlands; http://www.svi.nl/) to

exclude out-of-focus information. For the image deconvolution, default settings of

the programs were used except: (1) the “max iterations” was changed from 50 to 10

and (2) the “signal to noise ratio” was changed from 40 to 2. Deconvoluted images

were further processed for contrast or dark/light level manipulations using Adobe

Photoshop 5.0® software.

2.6. Protein expression analysis using denaturing gel

electrophoresis and Western blotting

Total proteins were isolated from the Agrobacterium infiltrated area of Nicotiana

leaves or transgenic Arabidopsis leaves expressing BiFC constructs. Approximately 15

mg of homogenised leaf tissues were incubated with 150 μl of 1 x sodium dodecyl

sulfate (SDS) sample buffer (62.5 mM Tris-HCl, 2% SDS, 5% β-mercaptoethanol, 10%

glycerol, 0.01% bromophenol blue, pH 6.8) at 95°C for 5 min. The suspension was

clarified by centrifugation at 16000 x g for 5 min. Aliquots (10 μl) of protein extract

were separated by duplicate 10% SDS-polyacrylamide gel electrophoresis (SDS-PAGE;

Laemmli, 1970). One gel was stained with Coomassie Brilliant Blue R250 to compare

http://www.svi.nl/


protein loading amounts while the proteins on the other gel were transferred to a

nitrocellulose membrane (Protran®, Schleicher and Schuell) using a semi-dry transfer

system (Bio-Rad) for immunoblotting analysis. Immunodetection of fusion proteins

was done with monoclonal anti-c-myc antibodies or anti-HA antibodies (Santa Cruz

Biotechnology Inc.) followed by Amersham ECL™ Western Blotting System (GE

Healthcare, Buckinghamshire, UK). Chemiluminescent signals were detected using

the Gel Doc™ XR and ChemiDoc™ XRS Gel Documentation System and the Quantity

One software (Bio-Rad). Images were then processed for contrast or dark/light level

manipulations using Adobe Photoshop 5.0® software.

Chapter 3. Establishment of a BiFC assay system in planta 42

Chapter 3

Establishment of a Bimolecular

Fluorescence Complementation

Assay System in planta


3.1. Introduction

As discussed in detail in Chapter 1, 14-3-3 proteins act as key regulators of a

large number of biological processes by binding as homo or heterodimers to target

proteins. In vitro, in silico and non-homologous in vivo systems have led to the

identification of more than 300 potential 14-3-3 target proteins (Roberts, 2003;

Mhawech, 2005; Schoonheim et al., 2007a; Chang et al., 2009; Paul et al., 2009).

However, it appears that such systems, as they do not truly represent the native

protein environment, can lead to conflicting or misleading results as demonstrated by

Athwal and co-workers when testing 14-3-3:nitrate reductase interactions using

different ion and pH conditions (Athwal et al., 2000). Additionally, as discussed

earlier in detail (Section 1.7), the high amino acid sequence similarities of 14-3-3

isoforms supports experimentally observed functional redundancy of 14-3-3s (van

Heusden et al., 1996; Rosenquist et al., 2000; Ferl et al., 2002; Alsterfjord et al., 2004)

whilst differences in binding affinities of 14-3-3 isoforms for a given target protein

suggest functional specificity (Bachmann et al., 1996b; Kanamaru et al., 1999;

Rosenquist et al., 2000; Sinnige et al., 2005). Hypothetically, an in vivo assay system

that allows for protein-protein interactions to be studied in the native environment

has the potential to resolve the above discrepancies. This could be used to study the

still open question of 14-3-3 isoform specificity during dimer formation or target

protein interaction.

Bimolecular fluorescence complementation (BiFC) allows for the visualisation of

protein-protein interaction in vivo (Hu et al., 2002). This technique requires that the

coding regions of two candidate proteins are transcriptionally fused to two

non-fluorescent fragments of YFP. Co-expression of interacting proteins results in

reconstitution of functional YFP, which can be detected by monitoring YFP

fluorescence, for example via fluorescence microscopy techniques. BiFC analysis was


first used to demonstrate protein-protein interactions in living mammalian cells (Hu et

al., 2002; Hu and Kerppola, 2003) and was subsequently adapted for the use in plant

systems (Bracha-Drori et al., 2004; Walter et al., 2004). These first publications

reporting the use of BiFC in plants demonstrated its successful application to the

analysis of protein interactions in transient systems using Nicotiana benthamiana

(Nicotiana) or Arabidopsis thaliana (Arabidopsis) plants or protoplasts. Hence,

visualisation of protein interactions between reported 14-3-3 target proteins and each

14-3-3 isoform in living cells using the BiFC analyses would allow the functional

specificity of 14-3-3 proteins to be explored and determined.

To perform such studies, a system has to be developed that allows for testing of

a large number of protein interactions in a time and material efficient way. This

requires cloning of cDNA sequences encoding for proteins to be studied into suitable,

easy to handle vectors. It also needs a reliable and robust analysis platform in planta

for the medium-throughput BiFC assays for 14-3-3 protien interactions.

The aims of this part of project were:

(1) Develop a T-DNA binary vector system suitable for BiFC analysis of transiently and

stably transformed plants.

(2) Generate a library of Arabidopsis 14-3-3 BiFC vectors for protein interaction

analysis.

(3) Establish and evaluate a test system for BiFC analysis of transiently and stably

transformed plants.


3.2. Results

3.2.1. Identification of a suitable vector system for a plant based BiFC assay

system

To adapt a BiFC assay system in planta for 14-3-3 interaction analyses, two sets

of BiFC expression vectors were obtained from Profs. Claudia Oecking, and Klaus

Harter, ZMBP, Pflanzenphysiologie, Universität Tübingen, Tübingen, Germany (Walter

et al., 2004). One set of vectors is designated as pUC-SPYNE and pUC-SPYCE (for

split-YFP N-terminal/C-terminal fragment expression), and the other is pSPYNE-35S

and pSPYCE-35S (Walter et al., 2004; Fig. 3-1). Both sets of vectors contain the same

BiFC cassettes, which allow for the insertion of the coding region and expression of

proteins of interest translationally fused to a protein tag and either a 155 amino acid

N-terminal (SPYNE) or an 84 amino acid C-terminal fragment (SPYCE) of the enhanced

yellow fluorescent protein (eYFP). The sequences encoding the fusion proteins are

expressed under the control of a constitutive cauliflower mosaic virus (CaMV) 35S

promoter and the nopaline synthase gene terminator (NosT; Fig. 3-1). The protein

tags, a c-myc epitope tag in SPYNE vectors and a hemagglutinin (HA) epitope tag in

SPYCE vectors, allow for the detection of fusion proteins using immunoblot analysis

via commercially available antibodies directed towards the protein tags.

The major difference between the two sets of vectors is that the pUC-SPYNE and

pUC-SPYCE vectors are based on pUC19 and are specially designed for transient plant

cell transformation (Walter et al., 2004). On the other hand, pSPYNE-35S and

pSPYCE-35S are T-DNA binary vectors suitable for Agrobacterium-mediated transient

or stable plant transformation (Walter et al., 2004). As it was anticipated that

14-3-3 proteins would be tested for interaction in both transiently and stably

transformed plants during this project, it was initially decided to use only the BiFC

vectors, pSPYNE-35S/pSPYCE-35S, which are based on T-DNA binary vectors.


Fig. 3-1. Schematic representation of expression cassettes in BiFC vectors (modified after

Walter et al., 2004)

The figure shows the basic features of BiFC expression cassettes in the vector pairs

pUC-SPYNE and pUC-SPYCE as well as SPYNE-35S and SPYCE 35S as described by Walter et al.

(2004). Genes coding for proteins of interest are cloned into the multiple cloning site (MCS) to

allow for translational fusions with a protein tag (c-myc: c-myc affinity tag; HA: hemagglutinin

affinity tag) and the N-terminal 155 amino acids of enhanced YFP (N-YFP or YN) or the 84

amino acids of the C-terminal end of eYFP (C-YFP or YC). The DNA and predicted amino acid

sequences and unique restriction sites of the MCS are shown at the top of the figure. The

fusion construct can be expressed under the transcriptional control of the CaMV 35S

promoter (35S), and the transcriptional terminator of the nopaline synthase gene from

Agrobacterium tumefaciens (NosT).

When transforming E.coli with these vectors for amplification, it was found that the

pUC-SPYNE and pUC-SPYCE were easily taken up by E. coli, but the pSPYNE-35S and

pSPYCE-35S were not (data not shown). A possible reason for this is the relatively

large size of the pSPYNE-35S and pSPYCE-35S vectors (> 13 kb), compared to that of

pUC-SPYNE and pUC-SPYCE (~4.8 kb). To improve the efficiency of the

transformation step and streamline it for the planned large-scale analyses of

protein-protein interactions, it was decided to transfer the BiFC cassettes into a high

copy vector of smaller size, which should also be suitable for transient and stable

transformation of plant cells.

pUC-SPYNE/SPYNE-35S

pUC-SPYCE/SPYCE-35S

XbaI HpaI StuI SpeI BamHI SalI XhoI KpnI SmaI

_|_______|____________|___________|_____|_________________|_____|_________|______|

TCTAGAGTTAACCGGGCTCAGGCCTGGCGCGCCACTAGTGGATCCATCGATAGTACTGTCGACCTCGAGGGTACCGCTCCCGGGATG

_S__R__V__N__R__A__Q__A__W__R__A__T__S__G__S__I__D__S__T__V__D__L__E__G__T__A__P__G__M

c-myc

35S MCS N-YFP (YN) NosT

HA

35S MCS C-YFP (YC) NosT


Binary vectors widely used for plant transformation have been summarised by

Hellens et al. (2000a). Many of these are larger than 10 kb and might have similar

drawbacks on genetic manipulation as described above for pSPYNE-35S/pSPYCE-35S.

Hellens et al. (2000a) also describe a vector series based on the high copy vector

pGreen, which has a relatively small size (3.3-7 kb). In addition, the pGreen series is

highly flexible with respect to components such as plant selectable markers,

promoters and terminators. This feature is a further advantage when transformation

of a plant with two or more different gene constructs is desirable (Hellens et al.,

2000b).

The pGreen vectors are widely used to generate transgenic plants via

Agrobacterium-mediated transformation (Hellens et al., 2000b; Vain et al., 2003; Cao

et al., 2007; Yang et al., 2008) and to perform transient gene expression in Nicotiana

leaves by agroinfiltration (Hamilton et al., 2002; Aparicio et al., 2005; Hellens et al.,

2005) or in onion epidermis by particle bombardment (Zhong et al., 2008). Based on

the versatility, copy number and transformation efficiency, it was decided to transfer

the BiFC cassettes from pUC-SPYNE/pUC-SPYCE into the pGreen vector backbone to

generate a new set of binary BiFC vectors for this study.

The pGreen vectors chosen as recipient vectors for the YN and YC cassettes were

pGreen0179 and pGreen0229. The two vectors contain different plant selectable

markers; pGreen0179 carries a hygromycin resistance gene (35S-hyg) whereas

pGreen0229 carries a bialaphos (Basta) herbicide resistance gene (Nos-bar; Fig. 3-2).

Using these two plasmids in combination is a convenient way of introducing two BiFC

constructs into the same plant as it allows for selection of two independent

transformation events, i.e. the transformation with two independent constructs

(Chapter 5).

Fig. 3-2. Generation of pGreen vector derivatives for the construction of BiFC vectors.

The MCS of pGreen0179 and pGreen0229 was modified by elimination of a number of restrictionsites, which were duplicated in the MCS of BiFC cassettes which were to be cloned into the pGreenvector backbones (Fig. 3-1). The first level of modification was done by eliminating the XhoI site toproduce the vectors pG179N and pG229N. The second level of modification was achieved byremoving the region between the SmaI and the SacI restriction sites to generate the vectorspG179NS and the pG229NS.Abbreviations: ColE1ori/pSa-ORI, E. coli/Agrobacterium origin of replications; LB/RB, T-DNAleft/right borders; Nos-bar/35S-hyg, plant Basta/hygromycin resistance genes; nptI, bacterialkanamycin resistance gene.


pG179N

MCS

pG229N

MCS

SacI digestionBlunted the protruding ends SmaI digestionReligation

EcoRV PstI BamHI

KpnI HindIII EcoRI SmaI SpeI XbaI NotI SacI| | | | | | | | | | |

GGTACCGGGCCCCCCCTCGACGGTATCGATAAGCTTGATATCGAATTCCTGCAGCCCGGGGGATCCACTAGTTCTAGAGCGGCCGCCACCGCGGTGGAGCTC

CCATGGCCCGGGGGGGAGCTGCCATAGCTATTCGAACTATAGCTTAAGGACGTCGGGCCCCCTAGGTGATCAAGATCTCGCCGGCGGTGGCGCCACCTCGAG

pG229NS

MCS

pG179NS

MCS

EcoRV PstI

KpnI HindIII EcoRI | | | | |

GGTACCGGGCCCCCCCTCGACGGTATCGATAAGCTTGATATCGAATTCCTGCAGCCCC

CCATGGCCCGGGGGGGAGCTGCCATAGCTATTCGAACTATAGCTTAAGGACGTCGGGG

pGreen0179

MCS

pGreen0229

MCS

XhoI / SalI double digestionReligation

EcoRV PstI BamHI

KpnI XhoI SalI HindIII EcoRI SmaI SpeI XbaI NotI SacI| | | | | | | | | | | | |

GGTACCGGGCCCCCCCTCGAGGTCGACGGTATCGATAAGCTTGATATCGAATTCCTGCAGCCCGGGGGATCCACTAGTTCTAGAGCGGCCGCCACCGCGGTGGAGCTC

CCATGGCCCGGGGGGGAGCTCCAGCTGCCATAGCTATTCGAACTATAGCTTAAGGACGTCGGGCCCCCTAGGTGATCAAGATCTCGCCGGCGGTGGCGCCACCTCGAG


The multiple cloning sites (MCS) of pGreen vectors are largely identical to the

MCS within the BiFC cassettes (Figs. 3-1, 3-2). Introducing the BiFC cassettes directly

into pGreen vectors would have created plasmids in which several of the MCS

restriction sites were present more than once, making the sites unsuitable for

subsequent cloning of genes of interest. To circumvent this problem, the pGreen

vectors were modified by deleting restriction sites in their MCS. The XhoI site in the

MCS of the pGreen vectors was eliminated by performing a digest using the enzymes

XhoI and SalI, followed by religation of the complementary ends generated (Fig. 3-2).

This resulted in the two vectors pG179N and pG229N. Furthermore, restriction sites

between SmaI and SacI were eliminated to generate the vectors pG179NS and

pG229NS. This left only a few duplicated restriction sites in the vector MCS and the

inserted BiFC cloning cassettes. The HindIII and EcoRI sites in these modified pGreen

vectors were maintained to allow for insertion of BiFC cloning cassettes (Fig. 3-2).

3.2.2. Construction of pGreen-based BiFC vectors

The BiFC cassettes were released from pUC-SPYNE and pUC-SPYCE by HindIII and

EcoRI double digest and inserted into the modified pGreen vectors, pG179N/pG229N

and pG179NS/pG229NS. The cloning strategy is depicted in Fig. 3-3 and resulted in

four pairs of BiFC vectors containing either 35S-hyg (pG179 derivatives) or Nos-bar

(pG229 derivatives) plant selection markers. These were designated pG179N-YN/-YC,

pG229N-YN/-YC, pG179NS-YN/-YC and pG229NS-YN/-YC (Fig. 3-3). Several unique

restriction sites suitable for subcloning DNA fragment of interest are present in these

vectors (Fig. 3-4). More specifically, the “N” series (pGxxxN-xx) vectors have two

unique cloning sites, HpaI and XhoI sites whilst the “NS” vector series (pGxxxNS-xx)

has more unique cloning sites (see Fig. 3-4 for details).

Fig. 3-3. Cloning strategy for the generation of pGreen-based BiFC vectors.

BiFC cassettes (35S::c-myc-YN or 35S::HA-YC) were isolated from pUC-SPYNE or pUC-SPYCE byHindIII/EcoRI digest and then inserted into HindIII/EcoRI linearised pGreen based vectorspG179N/pG229N and pG179NS/pG229NS. This resulted in two sets of pGreen-based BiFCvectors, carrying either the 35S-hyg or the Nos-bar plant selection genes.Abbreviations: bar/hyg, Basta herbicide/hygromycin resistance genes; c-myc/HA,c-myc/hemagglutinin epitope tags; N-YFP(YN)/C-YFP(YC), N/C-terminal fragment (amino acids1 to 155 or 156-239) of enhanced YFP; LB/RB, T-DNA left/right borders; MCS, multiple cloningsites; NosT, nopaline synthase gene terminator; 35S/Nos, cauliflower mosaic virus 35S/nopaline synthase gene promoters.

35S MCS NosTN-YFP (YN)

c-myc

pUC-SPYNE

SacI EcoRIHindIII

35S MCS NosTC-YFP (YC)

HA

pUC-SPYCE

SacI EcoRIHindIII

HindIII / EcoRIHindIII / EcoRI linearised

pG179N/pG229Nor

pG179NS/pG229NS


c-myc

pG179N-YN

SacI EcoRIHindIII

35S-hyg

RB

LB


HA

pG179N-YC

SacI EcoRIHindIII

RB

35S-hygLB


c-myc

pG229N-YN

SacI EcoRIHindIII

Nos-bar

RB

LB


HA

pG229N-YC

SacI EcoRIHindIII

RB

Nos-barLB


c-myc

pG179NS-YN

SacI EcoRIHindIII

35S-hyg

RB

LB


HA

pG179NS-YC

SacI EcoRIHindIII

RB

35S-hygLB


c-myc

pG229NS-YN

SacI EcoRIHindIII

Nos-bar

RB

LB


HA

pG229NS-YC

SacI EcoRIHindIII

RB

Nos-barLB


Fig. 3-4. Restriction maps of pGreen-based BiFC vectors.

Schematic representation of expression cassettes and polylinker sequences of thepG179N/pG229N derivative BiFC vectors (A) and the pG179NS/pG229NS derivative BiFCvectors (B). Unique restriction sites are indicated by an asterisk. Abbreviations are as indicatedin Fig. 3-3.

XbaI HpaI* SpeI BamHI SalI XhoI* SmaI

| | | | | | |


S R V N R A Q A W R A T S G S I D S T V D L E G T A P G M

35S MCS NosTYN or YCTag

EcoRV PstI BamHI

KpnI HindIII* EcoRI* SmaI SpeI XbaI NotI SacI

| | | | | | | | | | |

GGTACCGGGCCCCCCCTCGACGGTATCGATAAGCTTGATATCGAATTCCTGCAGCCCGGGGGATCCACTAGTTCTAGAGCGGCCGCCACCGCGGTGGAGCTC

pG179N-YN/-YCor

pG229N-YN/-YC

A

XbaI* HpaI* SpeI* BamHI* SalI XhoI* SmaI*

| | | | | | |


S R V N R A Q A W R A T S G S I D S T V D L E G T A P G M

35S MCS NosTYN or YCTag

KpnI HindIII* EcoRI* PstI

| | | |

GGTACCGGGCCCCCCCTCGACGGTATCGATAAGCTTGATATCGAATTCCTGCAGCCCC

pG179NS-YN/-YCor

pG229NS-YN/-YC

B



3.2.3. Cloning of Arabidopsis 14-3-3 open reading frames

The Arabidopsis 14-3-3 gene family is comprised of 15 genes that are called

General Regulatory Factor (GRF) with an Arabic number. Thirteen of these are

expressed while the remaining two are considered to be pseudogenes (Chevalier et al.,

2009). The expressed Arabidopsis 14-3-3 proteins are designated by Greek letters.

The nomenclature of the 14-3-3 family is summarised in Chapter 1 (Table 1-1).

In this study, a PCR approach was used to clone all ORFs of expressed

Arabidopsis 14-3-3 isoforms with the exception of 14-3-3 π (pi). The 14-3-3 pi was

not included because evidence for its transcription was not verified until after this

project was well underway (Sehnke et al., 2006; Cao et al., 2007). Primers for PCR

amplification of individual isoforms are listed in Table 2-1 (Section 2.1.2.4). To fuse

14-3-3 sequences to a protein tag and in frame with either the C- or N-terminal part

of the YFP coding region, an XhoI restriction enzyme site (CTCGAG) was introduced

into all 3’ primers. This also allowed the directional subcloning of 14-3-3

amplification products. This enzyme site was chosen as it eliminates the TAG stop

codon found in most of the 14-3-3 ORFs with minimal changes of surrounding base

pairs. The site was chosen also because all the Arabidopsis 14-3-3 cDNAs lack an

XhoI site. The 5’ primer included the ATG start codon of 14-3-3 ORFs and up to five

bases upstream of the ATG.

The N-terminal regions of 14-3-3s are essential for dimerisation, and truncation

of 14-3-3 ORFs by about 30 amino acids prevented dimerisation of these proteins

(Jaspert and Oecking, 2002; Walter et al., 2004). Thus, 14-3-3 sequences truncated

at their N-termini were used to assess non-specific interactions of 14-3-3 proteins in

negative control experiments. Non-specific interaction may occur due to molecular

crowding, leading to reconstitution of YFP without the requirement of target protein

interaction, a problem when using highly active promoters such as the CaMV 35S


promoter which leads to strong expression of proteins (Bhat et al., 2006; Lalonde et

al., 2008). To obtain these truncated 14-3-3s, a second set of primers was designed

for the 5’-ends. These primers were placed 78 to 99 bp downstream to the start

codon of the full-length ORF and included a second, in frame ATG present in all of the

14-3-3 ORFs. Amplicons obtained using these primers were expected to be

truncated between 26 and 33 amino acids at the N-terminal ends compared to the

full-length amplicons (Table 3-1). To differentiate between the two amplicons for

each of the 14-3-3 isoforms, full-length ORFs are indicated by an “X” after the isoform

name whilst the names of the truncated clones contain the symbol delta (Δ). For

example, the full-length ORF of the 14-3-3 isoform chi was named chiX whilst the

N-terminal truncated ORF of this isoform was named chiΔ (Table 3-1).

Plasmids containing either 14-3-3 cDNAs or cDNA libraries or self made cDNA

populations obtained from a mix of root and leaf mRNA were used as templates for

the PCR amplification of full-length and N-terminal shortened 14-3-3 ORFs as shown

in Table 3-1. The identities of all the PCR products were verified by sequence

analysis after subcloning (data not shown). In some cases nucleotide exchanges

were observed (Table 3-1). A representative result of PCR for amplification of the

14-3-3 ORFs from the leaf and root cDNA mixture is shown in Fig. 3-5. PCR products

were obtained for the full-length ORFs of the 14-3-3s kappa, nu and omega and for

the truncated ORFs of 14-3-3s chi, epsilon, mu and omega. These PCR products had

the expected amplicon size. The predicted size differences between full-length and

truncated versions of the omega isoform were observed (Fig. 3-5, lanes 8 and 9), and

are representative of those for other 14-3-3 isoforms (Fig. 3-5). Attempts to clone

14-3-3 iota ORFs using the leaf and root cDNA mix as template failed (Fig. 3-5, lanes 3

and 4). This is consistent with the reported observation that 14-3-3 iota is expressed

in Arabidopsis flowers but not in root and leaf tissues (Rosenquist et al., 2001).


Table 3-1. Templates and open reading frame specifications for cloned

Arabidopsis 14-3-3 sequences.

This table summarises the names of cloned 14-3-3 DNAs, the templates used to obtain these

DNAs with PCR-based cloning approaches and specifications of the 14-3-3 DNA sequences.

All the DNAs contain 14-3-3 open reading frames (ORFs) beginning with an ATG and ending

with the last amino acid encoding codon prior to the native stop codon. The stop codon was

omitted to allow translational fusion of 14-3-3 DNAs to other protein encoding DNAs.

Clones containing full-length 14-3-3 ORFs are designated with ‘X’ after the allele name.

Clones containing the shortened 14-3-3 ORFs designed for the deletion of the N-terminal

dimerisation domain are designated with ‘Δ’ after the allele name. The length of the

N-terminal deletions was determined by the position of a second in frame ATG start codon.

The details of the template sources are describe in the Materials and Methods. The cDNA

clones of 14-3-3 ORFs from the Arabidopsis stock centre are indicated by “*”. Nucleotide

sequence changes and resulting amino acid exchanges are shown in the table.

14-3-3 Template for PCR

amplification Sequence changes

(nucleotide/amino acid) Number of amino acids deleted

from the N-termini

chiX cDNA mix - -

chi∆ chiX - 32 aa

epsilonX cDNA library - -

epsilon∆ cDNA mix - 27 aa

kappaX CD3-360* - -

kappa∆ CD3-360* - 29 aa

lambdaX cDNA library C18G/silent mutation -

lambda∆ lambdaX - 29 aa

muX CD3-363* - -

mu∆ cDNA mix - 29 aa

nuX cDNA mix - -

omegaX CD3-202* A247G/Thr83Ala -

omega∆ cDNA mix - 27 aa

omicron∆ cDNA library - 27 aa

phiX cDNA mix - -

phi∆ phiX - 33 aa

psiX cDNA mix - -

psi∆ cDNA library - 26 aa

upsilonX CD3-203* C486T/silent mutation -

upsilon∆ CD3-203* C486T/silent mutation 29 aa


B

Lane 14-3-3

amplicon

Predicted

size (bp)

1 chiΔ 715

2 epsilonΔ 691

3 iotaX 819

4 iotaΔ 718

5 kappaX 759

6 muΔ 712

7 nuX 810

8 omegaX 792

9 omegaΔ 706

Fig. 3-5. PCR amplifications of Arabidopsis 14-3-3 encoding DNAs.

Several 14-3-3 encoding DNAs were PCR amplified using cDNA mixtures generated from

leaves and roots as templates and gene-specific primers for full-length and truncated 14-3-3

products. PCR products were analysed on a 1% agarose gel (A). The position of an 850 bp

molecular size marker is indicated. The identity and predicted sizes of the 14-3-3 amplicons

are shown in the table (B). Note, iotaX and iotaΔ were not amplified under the conditions

used in this experiment (see Section 3.2.3).

The amplicons of 14-3-3 ORFs, inserted into pCRII-TOPO cloning vectors, were

analysed for size and orientation in the vectors by restriction digestion. A

pCRII-TOPO construct can contain a 14-3-3 amplicon in either a forward orientation or

a reverse orientation. The orientation was defined by the relative position of the

5’-end of the 14-3-3 ORF to the unique HindIII site upstream the TOPO-TA cloning site

(Fig. 3-6 and pCRII-TOPO map, Invitrogen TA-cloning manual). If the 5’-end was

closer to the HindIII site, then the clone was designated as being in the forward

orientation; otherwise it was defined as being in the reverse orientation.

Fig. 3-6. Determining the orientation of cloned 14-3-3 encoding DNA fragments in TOPOvectors by restriction digestion.

Schematic diagrams demonstrate the elucidation of the orientation of cloned full-length 14-3-3phi coding sequences (phiX) in the pCRII-TOPO vector. The orientation of 14-3-3 fragments wasdefined by the relative position of the 5’-end of the 14-3-3 ORF to the unique HindIII site (asindicated by an asterisk) upstream of the TOPO-TA cloning site. If the 5’-end was closer to theHindIII site, then the clone was designated as being in the forward orientation (A), otherwisethe reverse orientation (B). The phiX contains an internal HindIII site that was used todifferentiate the orientation of positive clones by HindIII digestion. Plasmids isolated from twopositive clones were examined by HindIII digestion and 1% agarose gel electrophoresis (C). Theshorter HindIII fragment (393 bp) indicated the ‘forward’ orientation (lane 1); whilst the largerHindIII fragment (534 bp) indicated the ‘reverse’ orientation (lane 2). Additionally a bandrepresenting the vector backbone and the remainder of the cloned fragment was obtained ineach digest (4.39 kb in forward, 4.25 kb in reverse). M: 1 kb Plus DNA ladder (Invitrogen). Theposition of a 500 bp molecular size marker fragment is indicated.

C 1 2 M

500 bp

A 393 bp

B 534 bp

*

*



An example of an insert orientation analysis is shown in Fig. 3-6. The

orientation of the phiX ORF in pCRII-TOPO was determined by HindIII digestion and

agarose gel electrophoresis by taking advantage of two HindIII sites, one in the MCS of

the pCRII-TOPO vector and the other in the phiX ORF (Fig. 3-6 A-B). Depending on

the orientation, two different sets of HindIII fragment sizes were expected: the 393

bp/4.39 kb fragments with the forward orientation (Fig. 3-6 A and the lane 1 in C) and

the 534 bp/4.25 kb fragments with the reverse orientation (Fig. 3-6 B and the lane 2 in

C). All of the cloned 14-3-3 amplicons were analysed following this approach. To

allow for similar subcloning procedures, all clones used in subsequent experiments

were in the reverse orientation. The sequences of the inserts in these clones were

determined to confirm the presence of correct, non-mutated 14-3-3 ORFs. A

difference to the omega sequence At1g78300 as published in GeneBank was found in

the sequence of the omegaX clone. The nucleotide exchange from A to G at

nucleotide position 247 resulted in a deduced amino acid exchange from threonine to

alanine (Thr83Ala; Table 3-1). The omegaX ORF was PCR amplified using a

high-fidelity DNA polymerase (Section 2.2.5) from a cDNA clone obtained from the

Arabidopsis stock centre ABRC (clone No. CD3-202), so the nucleotide difference was

potentially in the original 14-3-3 omega cDNA clone.

3.2.4. Insertion of 14-3-3 ORFs into BiFC binary vectors

In general, cloned 14-3-3 ORFs were transferred from TOPO vectors into

pGreen based BiFC vectors generated during the work presented in this thesis. This

was achieved in most cases by EcoRV/XhoI digestion of the TOPO vectors to release

the 14-3-3 encoding DNA fragment and insertion of the 14-3-3 ORF into HpaI/XhoI

linearised pGreen-based BiFC vectors. This strategy allowed for the directional

cloning of 14-3-3 ORFs and translational fusion to protein tags and YFP fragments

contained in the BiFC vectors. Alternative restriction enzymes were used for the


subcloning of the 14-3-3 chi and lambda ORFs because an EcoRV site is present in

these two ORFs. In some cases, the PCR amplicons of 14-3-3 ORFs were directly

inserted into BiFC destination vectors to facilitate the cloning processes (in

circumvention of TOPO cloning). This was achieved by digesting the PCR products of

14-3-3 ORFs amplified with a high-fidelity Taq DNA polymerase with XhoI, a restriction

site introduced via the 3’-located amplification primer. The resulting blunt/XhoI

fragments were inserted into HpaI/XhoI linearised BiFC vectors. This strategy was

applied to the cloning of 14-3-3 kappaX into pG-229N-YN/-YC to generate the plasmid

pG-kappaXPCR-229N-YN/-YC (kappa-YN/-YC) as well as for the cloning of kappaΔ,

iotaX, iotaΔ, lambdaX, omegaX, omicronX, omicronΔ and upsilonΔ into the

pGreen-based BiFC vectors to generate the plasmids as shown in Table 3-2. In all

cases, the correctness of the 14-3-3 sequences inserted into destination vectors as

well as translational fusions to the protein tags and YFP fragments were confirmed by

sequence analysis (data not shown). Such sequence analyses indicated that

translational fusions of 14-3-3 full-length ORFs to both the YN and YC fragment were

correctly obtained for ten Arabidopsis 14-3-3 isoforms (chi, epsilon, iota, kappa, mu,

nu, omega, omicron, phi and psi) and in addition for the YN fusion of the full-length

14-3-3 lambda ORF (Table 3-2). Plasmids containing the upsilon isoform were not

used in subsequent work as errors in the coding region led to the loss of the ATG start

codon. In addition to full-length clones, correct YN and YC fusions were obtained for

seven N-terminally truncated 14-3-3s, namely epsilonΔ, kappaΔ, lambdaΔ, muΔ,

omegaΔ, phiΔ and psiΔ (Table 3-2).


Table 3-2. Plant expression vectors used in this study.

Characteristics of the plant expression vectors used in this study are shown, including the

35S-GFP containing pCAMBIA 1302 vector, the obtained BiFC cassettes and the generated

14-3-3-split YFP constructs in the pGreen vector backbones.

Plasmid name Description Bacterial selection

Plant selection

Source or reference

pCAMBIA 1302 35S-mGFP5-His x 6; 2x35S-hyg;

T-DNA-based binary vector Kanamycin Hygromycin CAMBIA

pUC-SPYNE 35S-c-myc-YN; pUC19-based Ampicillin - Walter et al., 2004

pUC-SPYCE 35S-HA-YC; pUC19-based Ampicillin - Walter et al., 2004

pG-chiX-179NS-YC 35S-chi-YC; 35S-hyg;

T-DNA-based binary vector Kanamycin Hygromycin This study

pG-chiX-179NS-YN 35S-chi-YN; 35S-hyg;


pG-epsilon∆-179N-YC 35S-epsilon∆-YC; 35S-hyg;


pG-epsilon∆-179N-YN 35S-epsilon∆-YN; 35S-hyg;


pG-epsilon∆-229N-YC 35S-epsilon∆-YC; pNos-bar;

T-DNA-based binary vector Kanamycin Basta This study

pG-epsilon∆-229N-YN 35S-epsilon∆-YN; pNos-bar;


pG-epsilonX-179N-YN 35S-epsilon-YN; 35S-hyg;


pG-epsilonX-229N-YC 35S-epsilon-YC; pNos-bar;


pG-epsilonX-229N-YN 35S-epsilon-YN; pNos-bar;


pG-iota∆PCR-229N-YC 35S-iota∆-YC; pNos-bar;


pG-iotaXPCR-229N-YC 35S-iota-YC; pNos-bar;


pG-iotaXPCR-229N-YN 35S-iota-YN; pNos-bar;


pG-kappa∆PCR-229N-YC 35S-kappa∆-YC; pNos-bar;


pG-kappa∆PCR-229N-YN 35S-kappa∆-YN; pNos-bar;


pG-kappaXPCR-229N-YC 35S-kappa-YC; pNos-bar;


pG-kappaXPCR-229N-YN 35S-kappa-YN; pNos-bar;


pG-lambda∆PCR-229N-YC 35S-lambda∆-YC; pNos-bar;


pG-lambda∆PCR-229N-YN 35S-lambda∆-YN; pNos-bar;


pG-lambdaX-179N-YN 35S-lambda-YN; 35S-hyg;

T-DNA-based binary vector Kanamycin Hygomycin This study


Table 3-2. Plant expression vectors used in this study (continued).

Plasmid name Description Bacterial selection

Plant selection

Source or reference

pG-mu∆-179NS-YC 35S-mu∆-YC; 35S-hyg;


pG-mu∆-179NS-YN 35S-mu∆-YN; 35S-hyg;


pG-muX-179NS-YC 35S-mu-YC; 35S-hyg;


pG-muX-179NS-YN 35S-mu-YN; 35S-hyg;


pG-nuX-229N-YC 35S-nu-YC; pNos-bar;


pG-nuX-229N-YN 35S-nu-YN; pNos-bar;


pG-omega∆-179N-YC 35S-omega∆-YC; 35S-hyg;


pG-omega∆-179N-YN 35S-omega∆-YN; 35S-hyg;


pG-omega∆-229N-YC 35S-omega∆-YC; pNos-bar;


pG-omega∆-229N-YN 35S-omega∆-YN; pNos-bar;


pG-omegaXPCR-179N-YN 35S-omega-YN; 35S-hyg;


pG-omegaXPCR-229N-YC 35S-omega-YC; pNos-bar;


pG-omicron∆PCR-229N-YN 35S-omicron∆-YN; pNos-bar;


pG-omicronXPCR-229N-YC 35S-omicron-YC; pNos-bar;


pG-omicronXPCR-229N-YN 35S-omicron-YN; pNos-bar;


pG-phi∆-179NS-YC 35S-phi∆-YC; 35S-hyg;


pG-phi∆-179NS-YN 35S-phi∆-YN; 35S-hyg;


pG-phiX-179NS-YC 35S-phi-YC; 35S-hyg;


pG-phiX-179NS-YN 35S-phi-YN; 35S-hyg;


pG-psi∆-179NS-YC 35S-psi∆-YC; 35S-hyg;


pG-psi∆-179NS-YN 35S-psi∆-YN; 35S-hyg;


pG-psiX-179NS-YC 35S-psi-YC; 35S-hyg;


pG-psiX-179NS-YN 35S-psi-YN; 35S-hyg;


pG-upsilon∆PCR-229N-YC 35S-upsilon∆-YC; pNos-bar;



3.2.5. Generation of an Agrobacterium library carrying pGreen-14-3-3-BiFC

constructs

The generated 14-3-3 full-length and N-terminally shortened 14-3-3 BiFC

plasmids were individually transferred into the Agrobacterium strain GV3101/pMP90

containing the pSOUP helper plasmid, which provides in trans activities required for

transformation of plants with pGreen based vectors (Hellens et al., 2000b). In this

library, each clone contained only one 14-3-3-split YFP construct, either the 14-3-3-YN

or the 14-3-3-YC. The presence of BiFC vectors in transgenic Agrobacteria was

confirmed by antibiotic selection and colony PCR analysis (Section 2.3.5). Examples

for the outcome of colony PCR analyses are shown in Fig. 3-7. In this experiment,

nine different Agrobacterium clones containing 14-3-3-split YFP plasmids were

analysed using a 14-3-3 isoform specific 5’-cloning primer (Table 2-1) as the forward

primer and a primer homologous to the coding region of YN (NYFPsp1; Table 2-2) or

YC (CYFPsp1; Table 2-2) as the reverse primer. Amplification products were analysed

by agarose gel electrophoresis. A single amplicon with the expected size was found

for each of the clones shown (Fig. 3-7).

3.2.6. Establishment of transient BiFC assay systems for 14-3-3 protein

interaction analysis

To establish the BiFC assay systems, a suitable plant system had to be identified.

In general, such a system could be based on transient or stable expression of BiFC

constructs. Agrobacterium-mediated transient transformation was first used as the

method for the delivery of transgenes into plant cells based on its simplicity, the

ability to obtain results rapidly when compared to stable transformation and the

potential for a medium-throughput evaluation of protein pair interactions.

35S 14-3-3X or 14-3-3Δ NosTYN or YCTag

5’ primer

3’ primer

A

1 2 3 4 5 6 7 8 9B

1 kb

Fig. 3-7. PCR based analysis of transgenic Agrobacterium lines containing 14-3-3-BiFCexpression vectors.

The presence of 14-3-3 BiFC expression vectors in transgenic Agrobacteria was confirmed viaPCR and gel electrophoresis. Cells picked from transgenic colonies were used directly in PCRsto provide the DNA template. PCRs were performed using 14-3-3 gene specific primers (A, 5’primer) in combination with either a N-YFP or C-YFP specific primer (A, 3’ primer) to confirmthe presence of the transgenes (see Materials and Methods for details). In the experimentshown, nine positive Agrobacterium clones were selected for the PCR based analysis. PCRproducts obtained from the corresponding constructs as listed in the table (C) were analyzedusing 1% agarose gel electrophoresis (B). The position of a 1 kb molecular size marker isindicated (B). The expected sizes of amplicons for each construct are summarised in (C).

Lane Construct Expected size (bp)

1 pG-epsilonX-229N-YN 961

2 pG-epsilonX-229N-YC 960

3 pG-epsilonX-179N-YN 961

4 pG-epsilon∆-229N-YC 877

5 pG-epsilon∆-179N-YN 878

6 pG-epsilon∆-229N-YN 878

7 pG-kappa∆PCR-229N-YN 854

8 pG-lambdaX-179N-YN 946

9 pG nuX-179N-YN 997

C



Agrobacterium infiltration (agroinfiltration) of leaves is a widely used transient

transformation method for gene function analysis, promoter analysis and protein

production (Voinnet et al., 2003; Wroblewski et al., 2005; Ueki et al., 2008). This

method was applied to many plant species and performed very well when used for

transient transformation of Nicotiana leaves (Romeis et al., 2001; Voinnet et al., 2003;

Witte et al., 2004; Hellens et al., 2005; Wroblewski et al., 2005). It was shown that this

method could also be used for the transient transformation of Arabidopsis thaliana

leaves (Bracha-Drori et al., 2004; Wroblewski et al., 2005). The stated advantages of

this method and its wide applicability for transient plant transformation led to the

decision to adapt this method for Nicotiana and Arabidopsis to study 14-3-3 protein

interaction.

3.2.6.1. Transient BiFC assay in Arabidopsis leaves using agroinfiltration

The experimental conditions described in Bracha-Drori et al. (2004) for transient

BiFC assays in Arabidopsis leaves were applied in this study where possible, including

the experimental procedures, the Agrobacterium strain (GV3101/pMP90) and the

Arabidopsis ecotype (Col-0). However, the present study differed from the published

work as different T-DNA binary vectors and a helper plasmid (pSOUP) and different

gene sets were used.

For transient transformation, two Agrobacterium strains, one harbouring a

14-3-3-YN and the other a 14-3-3-YC vector were co-infiltrated into the abaxial air

space of 4-5-week-old Arabidopsis leaves (Section 2.4.2). In an initial evaluation

experiment, non-treated leaves were used to detect background fluorescence, and

infiltration with buffer only or with only one 14-3-3 BiFC construct (e.g. epsilon-YN)

were used as negative controls. To evaluate the efficiency of the transgene delivery

method, a single Agrobacterium strain carrying the vector pCAMBIA 1302, which


encodes the GFP reporter gene under the regulatory control of a CaMV 35S promoter,

was used to infiltrate leaves (positive control).

Three days after infiltration, the lower epidermal layers of infiltrated leaves were

examined by fluorescence microscopy. Fluorescence was not detectable in

non-treated leaves (Fig. 3-8 A), in leaves infiltrated with infiltration buffer only (Fig.3-8

B), nor in leaves infiltrated with the 14-3-3 epsilon-YN construct only (Fig. 3-8 D).

GFP fluorescence was detected in only some of the leaves infiltrated with the 35S-GFP

clones (Fig. 3-8 C), suggesting the competency of Arabidopsis leaf cells for receiving or

expressing transgenes via the agroinfiltration method varied from plant to plant

and/or leaf to leaf.

Leaves co-infiltrated with a 1:1 volume ratio of Agrobacteria harbouring

epsilon-YN and epsilon-YC constructs showed fluorescence, which was stronger than

the background fluorescence observed in negative control experiments. This

suggested YFP reconstitution by 14-3-3 epsilon homodimer formation (Fig. 3-8 E).

For another 14-3-3 dimer combination, psi-YN and psi-YC, the BiFC fluorescence was

dim and could not easily be differentiated from background fluorescence (Fig. 3-8 F).

Notably, BiFC fluorescence by the epsilon or the psi homodimerisation was not

consistently observed in replicate infiltrations, or in a repeat experiment (data not

shown).

Strong background fluorescence was often observed in cells surrounding

wounded tissues of Arabidopsis (Fig. S1; Appendix II). The wounds were possibly

caused by the infiltration procedure. These wounded areas became yellowish three

days after infiltration, and demonstrated very strong, green background fluorescence

(Fig. S1). Such strong background fluorescence severely hindered the detection of

BiFC fluorescence driven by potential 14-3-3 dimer formation in adjacent cells, and

made the system impractical for studying protein-protein interactions.

Fig. 3-8. Bimolecular fluorescence complementation analysis of 14-3-3 dimerisation intransiently transformed Arabidopsis leaves.

Arabidopsis leaves (4-5-week-old) were: not infiltrated (no treatment control; A and A’) orinfiltrated with buffer only (infiltration control; B and B), an Agrobacterium strain harboring avector encoding a 14-3-3 epsilon-YN fusion protein (negative control; C and C’), anAgrobacterium strain harboring the 35S-GFP vector pCAMBIA1302 (positive control; D and D’),a mix of two Agrobacterium strains harboring vectors which encode either a 14-3-3 epsilon-YNor a 14-3-3 epsilon-YC fusion protein (epsilon BiFC; E and E’), a mix of two Agrobacteriumstrains harboring vectors which encode either a 14-3-3 psi-YN or a 14-3-3 psi-YC fusion protein(psi BiFC; F and F’). Fluorescence was monitored 3 days after infiltration using a fluorescencemicroscope (A to F) fitted with a GFP filter or under bright field (A’ to F’). GFP fluorescencewas clearly observable in leaves infiltrated with the positive control (D) and a low level of BiFCfluorescence was observed in leaves infiltrated with the two pairs of 14-3-3 BiFC constructs (Eand F), but fluorescence was absent or very low in control treatments (A to C). Scale bars = 50μm.

GFP filter Bright field

A

B

D D’

C

E

F

A’

E’B’

F’C’




Factors that affect the virulence of Agrobacterium or the physiological condition

of a plant would have an impact on the efficiency of transient assays (Wroblewski et

al., 2005). Wroblewski and co-workers examined several parameters affecting

transgene expression in the transient agroinfiltration assays, such as the density of the

Agrobacterium suspension, Agrobacterium strains, Arabidopsis ecotypes, effect of

culture conditions and chemical components (Wroblewski et al., 2005). Two of these

parameters were investigated as means of improving the BiFC assay system in this

study: these were the cell density of the Agrobacterium suspension and the

Arabidopsis ecotypes.

When the OD600 of the Agrobacterium cell suspension used for agroinfiltration

was increased from 0.2 to 0.6 or 1.0, the efficiency and reproducibility in GFP

transformation experiments increased slightly as observed by slightly stronger

fluorescence in more cells and a more uniform distribution of fluorescent cells (data

not shown). No significant difference was observed between infiltration

experiments in which cells at a density of OD600= 0.6 and 1.0 were used (data not

shown). In contrast to the positive GFP control, no improvement was observed in

14-3-3 co-infiltration assays (data not shown). Using other ecotypes of Arabidopsis

such as Col-6 and Wassilewskija-2 (WS-2), led to no obvious improvement in the

fluorescence intensity or the number of cells transformed (data not shown).

The results from these evaluation experiments suggested that agroinfiltration of

Arabidopsis leaves was not a very reliable technique to study 14-3-3 protein

interaction using BiFC, at least under the experimental conditions tested. Although

not all of the assay parameters were exhaustively examined, further optimisation of

this system would have delayed achieving the main aims of this project.

Consequently, other transient assay systems for BiFC analysis were tested.


3.2.6.2. Transient BiFC assay in Arabidopsis mesophyll protoplasts

To maintain the transient BiFC assay in the homologous system, i.e. test

Arabidopsis proteins in Arabidopsis, an alternative approach was needed to deliver

transgenes into Arabidopsis cells. Two other approaches widely used in transient

assays in Arabidopsis are biolistic bombardment and PEG-mediated transformation of

protoplasts (Ueki et al., 2008).

Microparticle bombardment is a widely used technique for transient gene

expression, and is used with a variety of plant species (Taylor and Fauquet, 2002), but

was not applied here as bombardment of Arabidopsis tissues is reportedly low in

efficiency and reproducibility (Helenius et al., 2000).

PEG-mediated transformation of protoplasts is an efficient method for transient

gene expression (Yoo et al., 2007), and has been used for in planta BiFC assays (Walter

et al., 2004; Schütze et al., 2009). Walter and co-workers demonstrated the

homodimerisation of a tobacco 14-3-3 isoform (T14-3c) in Arabidopsis cell culture

protoplasts using the BiFC assay (Walter et al., 2004). The feasibility of the

protoplast system was evaluated here using the established pGreen-based 14-3-3-split

YFP vectors described in this thesis (Section 3.2.4).

To maintain Arabidopsis cell suspension cultures as the protoplast source for the

transient assay is relatively time and labour consuming. An alternative source for

protoplasts is leaf mesophyll cells of soil-grown Arabidopsis plants (Kovtun et al., 2000;

Sheen, 2001; Yoo et al., 2007). High transfection efficiency (60%-90%) was reported

with the use of mesophyll protoplasts (Yoo et al., 2007). Thus it was decided to use

mesophyll protoplasts for a transient BiFC assay of 14-3-3 dimerisation.

To test the feasibility of the protoplast system for a transient BiFC assay,

approximately 2 x 104 mesophyll protoplasts were transfected with the constructs

indicated in Table 3-3. Protoplasts were tested for viability after transfection (data


not shown) and only viable protoplasts were analysed for fluorescence. In addition

to testing for fluorescence caused by GFP or YFP reconstitution, a filter set for the

fluorophore fluorescein isothiocyanate (FITC; Section 2.5.1) was used to visualise

autofluorescence of chlorophyll in the chloroplasts. Non-transformed protoplasts

showed no fluorescence with the GFP filter and only red chlorophyll autofluorescence

with the FITC filter (data not shown).

Following transfection with pCAMBIA 1302, a 35S-GFP construct used as a

positive transformation control, about 10% of the viable protoplasts (approximately

100 protoplasts) presented GFP fluorescence (Fig. 3-9 A, Table 3-3). The

fluorescence observed after transformation with the GFP construct was strong and

distributed throughout the cell, with a high intensity in the nucleus (Fig. 3-9 A, left).

In addition to GFP fluorescence, red chlorophyll autofluorescence was detected

indicating the position of chloroplasts (Fig. 3-9 A, middle).

BiFC fluorescence was observed using a GFP filter for all the tested 14-3-3

dimeric combinations. With 2 to 6.6 μg of total plasmid DNA used for the protoplast

transfection, the approximate frequency of protoplasts showing BiFC fluorescence for

each 14-3-3 dimeric combinations tested ranged from 0.1% to 5% (Table 3-3). There

was no significant correlation of the DNA amount used for transfection and the

frequency of protoplasts showing BiFC fluorescence from the tested 14-3-3 dimers.

BiFC fluorescence observed in protoplasts co-transfected with plasmids

pG-muX-179NS-YN and pG-muX-179NS-YC (muX-YN + muX-YC; Fig. 3-9 B) or

pG-psiX-179NS-YN and pG-psiX-179NS-YC (psi-YN + psi-YC; Fig. 3-9 C) indicated that

mu and psi homodimers accumulated in the cytosol and possibly in the nucleus (Fig.

3-9). A small degree of overlap between the BiFC and chlorophyll fluorescence

patterns was detected; consequently, it could not be unequivocally resolved if 14-3-3

psi-YN + psi-YCC

Fig. 3-9. Bimolecular fluorescence complementation analysis of 14-3-3 dimerisation intransiently transformed Arabidopsis mesophyll protoplasts.

Arabidopsis mesophyll protoplasts were transformed using a PEG based transformationprotocol with either a GFP expressing pCAMBIA 1302 vector (A) or co-transfected withconstructs encoding the 14-3-3 fusion proteins mu-YN and mu-YC (B) or psi-YN and psi-YC (C).Protoplasts were analysed 36 hours after transformation using a fluorescence microscopefitted with a GFP filter for the detection of fluorescence caused by GFP expression and YFPreconstitution upon 14-3-3 dimerisation (green), or a FITC filter for the simultaneousdetection of chlorophyll autofluorescence (red) and GFP/YFP fluorescence (green). Thebright field image is also shown for each cell. Scale bars = 50 μm.

GFP filter FITC filter Bright field

mu-YN + mu-YCB

GFPA



Table 3-3. Constructs used in the PEG-mediated transfection of Arabidopsis

mesophyll protoplasts for BiFC analysis of 14-3-3 dimerisation.

Highly purified pCAMBIA 1302 DNA (GFP positive control) or a combination of 14-3-3-BiFC

constructs was introduced into Arabidopsis leaf mesophyll protoplasts. The total amount of

plasmid DNA used is indicated. For BiFC experiments, both plasmids contributed equally to

the total amount of DNA used. The approximate frequency of protoplasts showing GFP or

BiFC fluorescence is shown.

Constructs Total plasmid DNA used

in transfections (μg)

Approximate frequency of

protoplasts showing fluorescence

pCAMBIA 1302

(GFP, positive control) 2.0 10.0%

chi-YN + kappa-YC 4.0 1.8%

chi-YN + nu-YC 4.0 1.4%

epsilon-YN + epsilon-YC 2.5 5.0%

iota-YN + iota-YC 6.0 4.0%

kappa-YN + kappa-YC 4.5 0.8%

mu-YN + mu-YC 3.0 3.0%

omega-YN + omega-YC 2.5 2.0%

omicron-YN + omicron-YC 3.4 0.1%

phi-YN + phi-YC 6.6 2.8%

psi-YN + psi-YC 2.0 3.0%

mu and psi homodimer formation occurred in chloroplasts (Fig. 3-9 B and C). From

these results, it appeared that the protoplast transfection system is suitable for the

evaluation of protein interactions using BiFC analysis. This method can potentially

be used to identify the subcellular localisation of such interactions. However, due to

the variation in fluorescent cell frequency, further improvement and optimisation of

this assay system appeared necessary.

3.2.6.3. Transient BiFC assay in Nicotiana leaves

Because of the limitations of the Arabidopsis agroinfiltration and protoplast

systems, which displayed low reproducibility and variable co-transfection frequency,


respectively, an alternative system, agroinfiltration of Nicotiana leaves, was assessed

for BiFC assays of 14-3-3 dimerisation. Nicotiana is used in a variety of transient

expression assays, including BiFC analyses for protein interactions (Bracha-Drori et al.,

2004; Walter et al., 2004; Desprez et al., 2007; Gampala et al., 2007; Waadt et al.,

2008).

Leaves of 5-7-week-old Nicotiana plants were infiltrated, through the abaxial air

spaces as described in Section 2.4.5, with a mixture of Agrobacterium clones

harbouring 14-3-3-YN and 14-3-3 YC constructs. In parallel, another set of leaves

was infiltrated with Agrobacteria harbouring the GFP-encoding vector pCAMBIA 1302

to serve as a positive control. As shown in Fig. 3-10 A, GFP fluorescence was

detected in the infiltrated leaf area, but only a few cells showed fluorescence.

Similarly in the leaves co-infiltrated with chi-YN and chi-YC constructs, the BiFC

fluorescence caused by homodimerisation of 14-3-3 chi isoform was detected only in

a small number of cells (Fig. 3-10 C). Further improvements in this approach were

needed for it to be a robust BiFC assay system.

It has been shown that high level of transgene expression following

agroinfiltration in Nicotiana leaves triggers post-transcriptional gene silencing (PTGS)

(Voinnet et al., 2003). Co-expression of a silencing suppressor, such as the p19

protein from tomato bushy stunt virus, with the proteins of interest in the

Agrobacterium-mediated transient assay system, can attenuate PTGS (Voinnet et al.,

2003). This strategy has been used in a number of transient assays to enhance

transgene expression (Hellens et al., 2005; Wroblewski et al., 2005; Jach et al., 2006;

Grefen et al., 2008), including BiFC analyses (Walter et al., 2004; Waadt et al., 2008;

Schütze et al., 2009).

Fig. 3-10. The impact of gene silencing in transiently transformed Nicotiana benthamianaleaves on bimolecular fluorescent complementation signals.

N. benthamiana leaves were infiltrated with Agrobacteria harboring the 35S-GFP vectorpCAMBIA 1302 (positive control, A and B; GFP) or with a mix of Agrobacteria harboringvectors expressing either a 14-3-3 chi-YN or a 14-3-3 chi-YC fusion proteins (C and D; chi-YN/chi-YC). These infiltrations were performed in the absence (A and C; -p19) or presence (Band D; +p19) of an Agrobacterium strain harboring a vector encoding the p19 silencingsuppressor. Injected leaves were monitored for epifluorescence, indicating GFP expression or14-3-3 dimerisation, 3 days after injection using a fluorescence microscope fitted with a GFPfilter (A to D, left). Bright field images of the infiltrated leaf areas are also shown (A to D, right).Scale bars = 50 μm.

GFP filter Bright field GFP filter Bright field

+ p19- p19

GFP

chi-YN/chi-YC

A

C

B

D



To determine if co-expression of BiFC constructs and the silencing suppressor

p19 could enhance transgene expression in the system used here, an Agrobacterium

strain containing the p19 construct was co-infiltrated with GFP or BiFC constructs into

Nicotiana leaves. As shown in Fig. 3-10, co-expression of GFP and p19 lead to a

strong increase in the number of cells showing GFP fluorescence as well as in the

intensity of the fluorescent signal observed in individual cells (Fig. 3-10 B). Similarly,

a strong increase in fluorescence was observed when Nicotiana leaves were

co-transformed with p19 and two 14-3-3 chi-split YFP constructs (chi-YN/chi-YC; Fig.

3-10 D). Approximately 90% of the cells in the infiltrated area demonstrated strong

fluorescent signals in both co-infiltration experiments. The fluorescence started to

appear 2-3 days post agroinfiltration and continued for up to 7 days. In general, the

best observation time for BiFC fluorescence was 4-5 days post-infiltration, after which

the fluorescent signal began to fade.

The results suggested that high expression of GFP and 14-3-3 proteins was

induced or stabilised by co-expression of p19 in Nicotiana leaf cells. The increase in

fluorescence intensity and number of cells showing fluorescence when using p19 was

reproducible and also observed using other 14-3-3 combinations (Fig. 3-11). This

improvement in signal output and experimental reliability was the reason the p19

silencing suppressor was included in subsequent transient BiFC analyses (Chapter 4).

3.2.6.4. Robustness of transient BiFC assays using Nicotiana leaves and

reproducibility of results

Overexpression of two proteins using highly active promoters, such as the CaMV

35S promoter, can lead to molecular crowding. In combination with ‘stickiness’ of

such proteins this may lead to false positive interactions, which could lead to

reconstitution of YFP and hence fluorescence. To verify that the observed BiFC


signals were specific to 14-3-3 dimerisation and not caused by auto-reconstitution of

the split YFP parts of the fusion proteins or stickiness of the expressed fusion proteins,

BiFC assays were performed using constructs encoding N-terminally truncated 14-3-3

proteins. Several in vivo and in vitro approaches have shown that such N-terminally

truncated 14-3-3s, lacking the dimerisation domains, are unable to dimerise (Wu et al.,

1997b; Abarca et al., 1999; Jaspert and Oecking, 2002; Walter et al., 2004). Thus,

such N-terminally truncated 14-3-3s could serve as negative controls for BiFC assays

(Walter et al., 2004). Any fluorescent signals obtained with such constructs would

reflect non-specific interactions of expressed proteins or tags.

The BiFC fluorescence of full-length 14-3-3 combinations was compared to that

of N-terminally truncated 14-3-3s to verify that only 14-3-3 dimerisation can lead to

reconstitution of YFP and hence fluorescence. Intense BiFC fluorescence was

observed in leaf cells infiltrated with full-length kappa-YN/kappa-YC, indicating the

formation of 14-3-3 kappa homodimers within the cells (Fig. 3-11, I-A). In contrast,

BiFC fluorescence was not detectable in leaves transformed with kappaΔ-YN/kappa-YC

or with kappaΔ-YN/kappaΔ-YC (Fig. 3-11, I-B and I-C, respectively). The results

confirmed that N-terminally truncated 14-3-3 kappa monomers were unable to

dimerise with a full-length or another truncated 14-3-3 kappa monomer, and that the

fluorescent signals were due to dimerisation of the full-length 14-3-3 proteins.

Western blot analysis was used to verify that lack of fluorescence was not

caused by a lack of expression of N-terminally truncated 14-3-3s. Protein extracts

were prepared from leaf areas infiltrated with full-length and N-terminally truncated

BiFC constructs. Proteins were separated by size using 10% denaturing

polyacrylamide gels and followed by immunoblot analyses using primary antibodies

directed against the c-myc or HA tags encoded in the BiFC constructs (Fig. 3-11, II).

Fig. 3.11. Bimolecular fluorescence complementation analysis of Arabidopsis 14-3-3 kappahomodimerisation and western blot analysis of the transgene expression in N.benthamiana leaves.

I. (A-C) Bright field images (upper panel) and epifluorescence images (lower panel) of thelower epidermal layer of leaves infiltrated with a mixture of Agrobacteriumsuspensions harbouring constructs encoding the 14-3-3 fusion proteins:kappa-YN/kappa-YC (A), kappaΔ-YN/kappa-YC (B) or kappaΔ-YN/kappaΔ-YC (C). Similaroutcomes were observed in duplicate experiments (data not shown). Scale bars = 50 μm.II. Immunodetection of 14-3-3 fusion protein expression with anti-c-myc (α-c-myc) and anti-hemagglutinin (α-HA) antibodies. Total proteins were extracted separately from the leavesshown in I, A-C, separated by 10% SDS-polyacrylamide gel electrophoresis and transferredto nitrocellulose membrane (lanes A-C, respectively). The membrane was probed with α-c-myc antibodies and then with α-HA antibodies as the primary antibodies. Horseradishperoxidase-conjugated anti-mouse antibodies were used as the second antibody. Labelledpolypeptides were visualised by the ECL™ Western Blotting System (GE Healthcare) and theGel Doc™ XR and ChemiDoc™ XRS Gel Documentation System (Bio-Rad).III. A duplicate gel to that shown in II stained with Coomassie Blue, showing protein amountsloaded. The abundant protein at about 54 kDa is likely the large subunit of ribulose-1,5-bisphosphate carboxylase/oxygenase (RubisCO).Positions of the 40 and 50 kDa protein markers are indicated. Note, irrelevant lanesbetween lanes B and C (in II and III) were removed.

Coomassie Blue staining

III

40

50

A B CkDa

A B CI


kappa-YN/kappa-YC kappaΔ-YN/kappa-YC kappaΔ-YN/kappaΔ-YC

Immunoblot

α-c-myc

α -HA

kDa

40

A B CII

50


A horseradish peroxidase-conjugated anti-mouse antibody was used as secondary

antibody, allowing for the simultaneous visualisation of YN and YC fused 14-3-3s.

Two intensely labelled bands with the expected sizes, i.e. 47 kDa for kappa-YN and 44

kDa for kappa-YC, were detected in extracts from leaves infiltrated with full-length

14-3-3 kappa constructs (Fig. 3-11, II lane A). Two bands of 39 kDa and 36 kDa were

observed in extracts obtained from leaves infiltrated with the N-terminally shortened

14-3-3 kappa constructs (Fig. 3-11, II lane C). These sizes corresponded to the

expected reduction in molecular weight of the 14-3-3s caused by the loss of 29 amino

acids at the N-terminal end of 14-3-3 kappa.

Other full-length and N-terminally truncated 14-3-3s were tested in the BiFC

assay as described above to confirm that the results obtained for the two forms of

14-3-3 kappa could be generalised for 14-3-3s (Table 3-4). As with N-terminally

truncated 14-3-3 kappa, none of the tested dimeric combinations containing at least

one truncated 14-3-3 showed intense BiFC fluorescence when compared to full-length

14-3-3s. Most of the combination with one or two truncated 14-3-3s displayed no

detectable BiFC fluorescence at all, whilst a few showed weak fluorescence in a small

number of cells (low level; Table 3-4 and data not shown). These data supported the

notion that BiFC fluorescence was caused by 14-3-3 dimerisations and not because of

YFP reconstitution by other mechanisms or to a general ‘stickiness’ of the proteins.

The results further supported that of others showing N-terminally truncated 14-3-3s

can serve as negative controls in 14-3-3 dimerisation experiments using BiFC

(Chapters 4 and 5).

3.2.7. Identification of subcellular distribution of 14-3-3-BiFC dimeric

complexes

When observing BiFC fluorescence from 14-3-3 dimerisations in intact leaves,


Table 3-4. Negative controls of 14-3-3 dimerisation tested in the transient

BiFC assay in Nicotiana leaves.

Selected N-terminally truncated 14-3-3 mutants were used to test for 14-3-3 dimerisations in

the transient BiFC assay. Some dimeric combinations showed undetectable BiFC

fluorescence and others showed low levels of fluorescence.

Constructs BiFC*

epsilonΔ-YN/epsilonΔ-YC ND

iota-YN/iotaΔ-YC Low level

kappa-YN/iotaΔ-YC Low level

kappa-YN/kappaΔ-YC ND

kappaΔ-YN/kappa-YC ND (Fig. 3-11)

kappaΔ-YN/kappaΔ-YC ND (Fig. 3-11)

muΔ-YN/chi-YC ND

mu-YN/muΔ-YC Low level

muΔ-YN/mu-YC ND

muΔ-YN/muΔ-YC Low level

omegaΔ-YN/omega-YC Low level

omegaΔ-YN/omegaΔ-YC Low level

* ND: not detectable. Low level: few cells showed weak BiFC

fluorescence in the infiltrated leaf areas.

for instance the kappa homodimerisation shown in Figures 3-11 A and 3-12 A, 14-3-3

dimers were usually detectable in the cytoplasm and sometimes also in nucleus-like

structures (Fig. 3-12 A). However, it was often difficult to identify subcellular

structures, such as the nucleus, with absolute certainty. To improve the resolution

and quality of the observation and imaging of fluorescent signals, peels of the lower

leaf epidermis were used. This procedure reduced the number of layers of cells for

microscopic observation, which dramatically increased image resolution (Fig. 3-12 C

and D). This led to more confident identification of the subcellular localisation of

14-3-3-BiFC dimeric complexes. In addition, this approach also allowed for an

Fig. 3-12. Comparison of 14-3-3 dimerisation mediated bimolecular fluorescencecomplementation signals in lower epidermal cells of intact leaves and in cells of peeledepidermal layers of Nicotiana benthamiana.

14-3-3 dimerisation, as indicated by reconstitution of YFP, was detected in N. benthamianaleaves infiltrated with two Agrobacterium strains, one carrying an expression construct for14-3-3 kappa-YN and the other a 14-3-3 kappa-YC expression vector. Epifluorescence imageswere taken from the lower epidermal layer of intact leaves (A) or from lower epidermalpeels (C). The fluorescent images demonstrate the increase in resolution of labeledintracellular structures such as the nucleus when using peeled epidermal layers (C) overintact leaves (A). In peeled epidermal cells (C), 14-3-3 kappa dimerisation was clearlydetectable via YFP fluorescence in the cytosol (Cyt) and the nucleus (N). In contrast, theBiFC signals of 14-3-3 kappa dimerisation were very low or below detectable levels in thenucleolus (No), in the vacuolar space (V) or in chloroplasts (Chl) surrounding the nucleus.Corresponding bright field images are shown (B and D). Scale bars = 50 μm.


A

C

B

D

N

V

Chl

Cyt

N

V

Cyt

No



estimation of fluorescence intensity caused by dimerisation in the different

subcellular compartments. As shown in Fig. 3-12 C, the fluorescence intensity of

14-3-3 kappa homodimerisation in nucleus-like structures was much stronger than

that observed in the cytoplasm, suggesting that kappa homodimers accumulated to

relatively high levels in the nucleus. A darker, possibly un-labelled structure

observed within the labelled nucleus resembled a nucleolus, suggesting the 14-3-3

dimer were low or absent from the nucleolus (Fig. 3-12 C and Fig. 3-13 A).

Fig. 3-13. Identification of the subcellular distribution of 14-3-3-BiFC dimeric complexes in

transiently transformed N. benthamiana leaves.

Epifluorescence images were obtained from abaxial epidermal peels of N. benthamiana

leaves infiltrated with a mixture of Agrobacterium suspensions harbouring constructs

encoding kappa-YN/kappa-YC (A-D) or phi-YN/omega-YC (E-H). The epidermal peels were

stained with 4’, 6-diamidino-2-phenylindole (DAPI) to detect the nuclei. BiFC fluorescence

from 14-3-3 dimerisations was detected as green under filters for GFP (A and E) and FITC (B

and F). Chlorophyll autofluorescence was red under the FITC filter (B and F). DAPI-stained

nuclei presented blue-white fluorescence under the DAPI filter (C and G). The kappa

homodimer and the phi-omega heterodimer localised to the cytosol, the nuclei (N) but were

absent from the nucleoli (No) and chloroplasts (Chl). DAPI-staining outside the cell was due to

debris. Scale bars = 50 μm.


The nuclear-like structure was confirmed to be the nucleus by 4’,

6-diamidino-2-phenylindole (DAPI) staining. Epidermal cells showing BiFC

fluorescence due to kappa homodimerisation (Fig. 3-13 A) or phi-omega

heterodimerisation (Fig. 3-13 E) were co-stained with DAPI, a fluorescent dye that

binds to double-stranded DNA resulting in blue-white fluorescence when excited with

UV light (Kubista et al., 1987). Using an appropriate filter set for DAPI (Section 2.5.1),

the location of the nuclei was confirmed to coincide with the observed nuclear-like

structures exhibiting BiFC fluorescence (Fig. 3-13). Chlorophyll autofluorescence

detectable as red fluorescence when using a FITC filter (Fig. 3-13 B and F) was used to

identify the location of chloroplasts. By comparison of FITC filter images with BiFC

images and overlays of the two, it was shown that both, kappa homodimers and

phi-omega heterodimers were absent from chloroplasts (Fig. 3-13).

A negative impact of using epidermal peels was the dramatic decrease of the

number of cells showing fluorescence when compared to intact leaves. After peeling,

approximately 5% to 15% of the epidermal cells still remained fluorescent. This

decrease may have been largely due to wound effects caused by the invasive

preparation of epidermal peels and by removing and observing only one cell layer.

Despite this reduction in fluorescent cell numbers, sufficient cells were available for

observation and determination of BiFC distribution. Thus, it was considered that the

improvement in image quality and the enhanced ability to clearly identify subcellular

localisation of 14-3-3 dimers outweighed the reduction in fluorescent cell numbers.

Thus, epidermal peels were used for all subsequent 14-3-3 dimerisations tested

(Chapter 4).


3.3. Discussion

3.3.1. Generation of pGreen-based BiFC vectors

A series of pGreen-based BiFC vectors was generated, which can be used to test

any two proteins of interest for interaction in living plant cells (Section 3.2.2). In

contrast to previously described vectors, the plasmids developed here are based on

the relative small, high copy number pGreen plasmid, which is easy to handle when

preparing plasmid DNA and in bacterial transformation experiments. In addition, the

flexibility of the pGreen series given by the large number of available cloning cassettes

(Hellens et al., 2000b), is maintained in the BiFC vectors produced in this study,

allowing for the use of various selection markers, promoters and terminators.

A library of Arabidopsis 14-3-3 ORFs in the generated BiFC vectors was

established, including eleven out of the thirteen expressed Arabidopsis 14-3-3

isoforms were inserted into the YN vectors and ten out of the thirteen were cloned

into the YC vectors (Table 3-2). These constructs comprise a basic 14-3-3-split-YFP

library that can be used to analyse localisation and specificity of 14-3-3 dimerisation.

Furthermore, this library can be used to test putative 14-3-3 target proteins for

interaction with 14-3-3 isoforms (Chapters 4). Moreover, a set of 14-3-3

N-terminally shortened ORFs were inserted into pairs of BiFC YN and YC vectors.

These can be used to study the impact of a loss of the dimerisation domain on 14-3-3

dimer formation and on target protein interaction. The experiments described in

this chapter (Section 3.2.6.4) clearly show that such truncated versions do not interact

with each other or with a full-length 14-3-3 protein, thus confirming their use as a

suitable negative control for molecular crowding or non-specific protein interaction as

shown previously by others (Walter et al., 2004).


3.3.2. The suitability of Arabidopsis mesophyll protoplasts for transient

BiFC assays

Three transient assay systems, Arabidopsis mesophyll protoplast transfection,

agroinfiltration of Arabidopsis leaves and agroinfiltration of Nicotiana leaves were

tested for their suitability in studies of protein interactions using BiFC. BiFC

fluorescence indicating 14-3-3 dimerisation was observed in Arabidopsis mesophyll

protoplasts chemically transfected with two pGreen-based 14-3-3-BiFC plasmids

(14-3-3-YN and 14-3-3-YC; Fig. 3-9). Ten pairs of 14-3-3 dimers have been tested in

this assay system and all of them showed BiFC fluorescence (Table 3-3).

Generally, each PEG-mediated protoplast transfection involves combining 10-20

μg of total plasmid DNA (10 μl) with 2 x 104 protoplasts (100 μl), although the plasmid

DNA/protoplast ratio for the transfection should be examined empirically (Yoo et al.,

2007). Following this general procedure, a transformed protoplasts frequency of

90% can be expected (Yoo et al., 2007). In the experiments conducted here, the

frequency of protoplasts successfully transformed was below the expected value.

For example, the use of 2 μg of a single 35S-GFP carrying plasmid for transfection

resulted in approximately 10% of the protoplasts exhibiting GFP fluorescence (Table

3-3). Co-transfection of two 14-3-3-BiFC constructs using 2 μg to 6.6 μg of total

plasmid DNA in a 1:1 ratio led to an even lower frequency of fluorescent cells

(approximately 0.1% to 5%; Table 3-3). The lower frequency of fluorescent cells in

BiFC experiments compared to GFP experiments was expected as BiFC fluorescence

requires successful co-transformation and co-expression of two BiFC constructs whilst

fluorescence in GFP transfection experiments requires only one plasmid to be

successfully taken up and expressed by the cells.

The overall low transformation rate indicated this protoplast system has limited

value for mass screening of protein interactions using BiFC. Instead, a protoplast


approach may be useful for studies investigating a small number of proteins.

Furthermore, this technique may be useful when transfecting protoplasts derived

from transgenic plants already expressing one of the BiFC constructs.

3.3.3. Assessment of BiFC assay using transient expression in Arabidopsis

and Nicotiana leaves via infiltration with Agrobacteria

Agrobacterium infiltration (agroinfiltration) is widely used to deliver transgenes

into plants for transient and stable analyses of gene function, promoter activity and

for protein interaction studies. This approach can be used for analysis in

homologous and non-homologous plant systems. The present study initially

favoured performing transient BiFC assays of Arabidopsis 14-3-3 isoforms in the

homologous Arabidopsis system using agroinfiltration. This approach has been

previously used to demonstrate interactions of α and β subunits of the Arabidopsis

farnesyltransferase (Bracha-Drori et al., 2004).

To evaluate the agroinfiltration of Arabidopsis leaves for the BiFC analysis of

14-3-3 dimerisations, as many of the parameters as possible used by Bracha-Drori et

al. (2004) were maintained in the present study, including the Agrobacterium strain

(GV3101/pMP90), Arabidopsis ecotype (Col-0) and experimental conditions.

Differences were the T-DNA binary vectors used (pGreen-based instead of

pCAMBIA-based) and the proteins of interest tested (14-3-3 proteins instead of

farmesyltransferase subunits). The results obtained with 14-3-3 proteins

demonstrated a high variability of fluorescence intensity and cell numbers showing

BiFC fluorescence (Fig. 3-8 E and F and data not shown). From these results, it was

difficult to determine which step of the assay caused this problem. Possible reasons

were that the transgenes were not efficiently delivered into a large number of plant


cells, that the transgenes were not well expressed, and/or that the 14-3-3 isoform did

not dimerise.

A successful agroinfiltration based BiFC assay requires that transgenes are

delivered into and expressed by the cells to a detectable level. For the BiFC assay, an

additional requirement, which must be fulfilled, is that the co-expressed candidate

proteins (here 14-3-3 proteins) interact with each other in a way that allows the YN

and YC parts of the YFP protein to interact, leading to BiFC fluorescence. The

efficiency of the gene delivery method can be evaluated, at least for the delivery of

one gene construct, using a 35S-GFP construct. A commercially available

GFP-containing vector, pCAMBIA 1302, was used for this purpose during this study.

This vector was chosen as it contained the same pCAMBIA 1302 backbone as the BiFC

vectors used by Bracha-Drori et al. (2004). This evaluation method confirmed the

low reproducibility of the gene delivery method (data not shown). To determine

whether the vector backbone had an impact on the frequency of transformation

events, another GFP construct, the pGreen-based 35S-GFP plasmid, was tested. The

use of this plasmid did not result in transformation events (data not shown). In

addition, agroinfiltration of Arabidopsis leaves and the subsequent 14-3-3 BiFC assays

did not generate reproducible results (Fig. 3-8 and data not shown). Taken together,

the results suggested that a) the nature of the Ti binary vector may influence the

performance of agroinfiltration of Arabidopsis leaves, b) to achieve high

transformation efficiency and high reproducibility of results would require elaborate

optimisation procedures to be performed.

Since two living organisms, plant and bacterium, participate in the process of

Agrobacterium-mediated transient assays, factors that affect the virulence of

Agrobacteria or the physiological condition of plants could have an impact on the

efficiency of agroinfiltration (Wroblewski et al., 2005). Using β-glucuronidase (GUS)


activity as a reporter, Wroblewski and co-workers (2005) examined several parameters

affecting transformation performance in these transient assays. The parameters

were evaluated such as the efficiency of several Agrobacterium strains, including the

strain GV3101 used in the present study, on four plant species: lettuce, tomato, N.

benthamiana and Arabidopsis (Col-0). Arabidopsis showed the least GUS activity

among the tested plant species, no matter which Agrobacterium strain was used

(Wroblewski et al., 2005). Additionally, transient GUS expression differed between

Arabidopsis ecotypes. A higher level of GUS expression was observed in the ecotype

WS than in Col-0 (Wroblewski et al., 2005). Wroblewski and co-workers (2005)

further investigated the effect of several viral silencing suppressors on inhibition of

PTGS to increase the level of transient expression in Arabidopsis. However, none of

the tested silencing suppressors, including p19, had a detectable effect on transient

GUS expression in Col-0. In the present study, attempts to fine-tune some of the

factors mentioned above to improve the outcome and reproducibility of transient

14-3-3 BiFC assay in Arabidopsis leaves (such as Arabidopsis ecotypes and p19) were

not successful (data not shown).

The suitability of Nicotiana leaf infiltration was investigated in parallel to the

study using Arabidopsis. The Nicotiana system showed higher transformation

efficiencies, stronger signals and could be optimised to demonstrate 14-3-3

dimerisation with great reproducibility. Combining the BiFC results obtained using

Nicotiana and Arabidopsis, the results presented here supported the findings of

Wroblewski et al. (2005) that the difference between plant types with respect to

transgene expression following agroinfiltration can be significant. Higher levels of

transgene expression following agroinfiltration of leaves were found for Nicotiana

than for two of the Arabidopsis ecotypes tested in the present study. Moreover, a

pGreen-based 35S-GFP construct was successfully introduced into Nicotiana leaves by


agroinfiltration but not into Arabidopsis leaves (data not shown). Furthermore,

when introducing a PTGS suppressor, p19, into the BiFC assay system, a strong

improvement of fluorescence, indicating improved or more stable transgene

expression was observed in Nicotiana leaves (Fig. 3-10), but not in Arabidopsis Col-0

(data not shown).

In summary, the Nicotiana-based BiFC assay system, as optimised in this study,

can serve as a robust, reliable and easy to perform assay system suitable for the

investigation of 14-3-3 protein interactions. As such, this system was employed in

further studies of 14-3-3 dimerisation and to determine the localisations of 14-3-3

dimers (Chapter 4).

Chapter 4. Visualisation of 14-3-3 dimerisation in living plant cells 87

Chapter 4

A Systematic Survey of

Subcellular Dimerisation of

Arabidopsis 14-3-3 Isoforms by

Bimolecular Fluorescence

Complementation


4.1. Introduction

4.1.1. Dimerisation of 14-3-3 proteins

14-3-3 proteins are highly conserved eukaryotic regulators coordinating a

variety of biological processes by physically interacting with phosphorylated target

proteins (Roberts, 2003). Native 14-3-3 proteins are always found as dimers (Aitken,

1996). In general these can be either homodimers or heterodimers.

Immunoprecipitation experiments followed by western blot analyses using 14-3-3

isoform specific antibodies indicated that different mammalian 14-3-3 isoforms show

differences in homo- or heterodimer formation. For example, the human 14-3-3

sigma isoform preferentially formed homodimers (Wilker et al., 2005), whilst the

epsilon isoform appeared to form exclusively heterodimers (Chaudhri et al., 2003).

The gamma isoform was able to homodimerise and to heterodimerise with the 14-3-3

isoforms beta, epsilon, zeta and eta (Chaudhri et al., 2003). A preference for

heterodimerisation was also reported for the two yeast 14-3-3 isoforms BMH1 and

BMH2 (Chaudhri et al., 2003). In plants, five recombinant Arabidopsis 14-3-3

isoforms (chi, psi, phi, omega and upsilon) were demonstrated to dimerise with each

other using yeast two hybrid assays or in vitro dimerisation assays combined with

native PAGE analyses (Wu et al., 1997b). In addition, recombinant Arabidopsis

14-3-3 psi and lambda were shown to form both homodimers and heterodimers in

vitro (Abarca et al., 1999). From these results, it appears that unlike those of

mammals or yeast, plant 14-3-3 isoforms do not show any preference for homo- or

heterodimer formation.

Structural and biochemical evidence suggest that the N-terminal domain of

14-3-3 monomers is required for dimerisation. X-ray crystal structure analysis of the

human 14-3-3 zeta isoform indicated that the N-terminal helix α1 of one monomer

interacts with helices α3 and α4 of the other monomer to form the dimerisation


interface (Liu et al., 1995). Further interaction studies performed on Arabidopsis

14-3-3 isoforms using in vitro dimerisation and yeast two hybrid assays showed that

the helices α1 to α4 are sufficient for dimerisation and that deletion of helix α4 leads

to dimerisation failure (Wu et al., 1997b; Abarca et al., 1999). Truncation studies of

the human 14-3-3 zeta isoform also demonstrated that an N-terminal truncated

mutant form was unable to dimerise but was still able to bind to its target protein

(Luo et al., 1995). In addition to verifying the role of the N-terminus in dimerisation,

Luo et al. (1995) also demonstrated that the C-terminal domain of 14-3-3 proteins

may contribute to binding of target proteins.

The dimeric structure of 14-3-3 proteins is related to their functions. A

monomeric form of the N-terminally truncated, human 14-3-3 zeta, although it can

bind to its target protein Raf-1, is unable to activate the target protein (Tzivion et al.,

1998). This strongly suggests that 14-3-3 dimerisation is important to at least part of

14-3-3 function. Additionally, it is widely proposed that 14-3-3 dimers act as

adaptors or scaffold proteins which bring two distinct target proteins together,

thereby requiring more than one target binding site (Tzivion et al., 2001; Roberts,

2003; Chevalier et al., 2009). Furthermore it appears that 14-3-3s, when binding to a

single target protein, do so by interacting with multiple binding sites within the target

protein. Both modes of action imply that 14-3-3 proteins must act as dimers. The

number of potential dimer combinations could also account for the large number of

experimentally verified and predicted target proteins if one assumes that 14-3-3

dimers have some kind of target specificity. Thus, it can be postulated that different

14-3-3 dimeric combinations may be related to different roles of 14-3-3s in cells.

So far, only a selection of Arabidopsis 14-3-3s has been tested for dimerisation

using in vitro approaches or yeast two hybrid analyses (Wu et al., 1997b; Abarca et al.,

1999). It remains elusive whether all members of the Arabidopsis 14-3-3 protein


family are able to freely dimerise with each other, especially in the native plant system.

Homodimerisation of a tobacco 14-3-3 isoform (T14-3c) using transient BiFC analysis

is the only direct evidence to date demonstrating 14-3-3 dimerisation in living plant

cells (Walter et al., 2004). Preferences for homo- and/or heterodimer formation of

14-3-3 proteins, as observed in animal systems, have not been investigated in plants

so far. This study will investigate whether Arabidopsis 14-3-3 isoforms have any

preference for dimer formation in vivo.

4.1.2. Subcellular localisation of 14-3-3 proteins in plants

In general, 14-3-3s are distributed widely throughout the cell and are usually

regarded as cytosolic proteins (Ferl et al., 2002; Alsterfjord et al., 2004). However, a

variety of studies in plants have demonstrated other intracellular distributions of

14-3-3 proteins. Immunohistochemical analyses showed that Arabidopsis 14-3-3

proteins are located in the nucleus (Bihn et al., 1997) and in chloroplasts (Sehnke et al.,

2000). Immunoblotting analyses using isoform specific antibodies confirmed this by

demonstrating the presence of the 14-3-3 isoforms epsilon, mu, nu and upsilon in

chloroplastic protein fractions (Sehnke et al., 2000). The same authors also

demonstrated the absence of 14-3-3 omega from chloroplastic fractions, thereby

showing isoform specific chloroplast localisation (Sehnke et al., 2000). In barley,

14-3-3 proteins were found in the matrix protein fraction of isolated mitochondria

using immunoblotting analyses (Bunney et al., 2001). Potential target proteins for

14-3-3 action in both, chloroplasts and mitochondria, are the organelle specific ATP

synthases (Bunney et al., 2001). In addition to immunological approaches, a number

of proteomics and GFP fusion studies revealed the diversity of subcellular localisation

of 14-3-3 proteins in Arabidopsis. The World Wide Web based SUBcellular protein

location database for Arabidopsis proteins (SUBA, http://www.suba.bcs.uwa.edu.au;

Heazlewood et al., 2007) summarises and compiles data from predicted and

http://www.suba.bcs.uwa.edu.au/


experimentally documented subcellular localisations for most Arabidopsis proteins.

This database was used here to establish the current view of subcellular localisations

of Arabidopsis 14-3-3 isoforms (Section 4.2.1).

Of the Arabidopsis 14-3-3 protein family, five members (chi, psi, phi, omega and

upsilon) have been shown to dimerise with each other in vitro (Wu et al., 1997b). Of

those, only psi, phi and omega have similar subcellular locations as revealed using

14-3-3-GFP fusion protein approaches or proteomic analyses (Table 4-1). The lack of

data with regards to 14-3-3 dimer formation and subcellular localisation justifies a

systematic in planta study aimed at testing the current knowledge, which is based

largely on in vitro studies, and to extend analyses to a larger number of 14-3-3

isoforms. The advantage of such a systematic study would be its comprehensiveness.

A single experimental procedure with defined conditions would greatly contribute to

comparability of data, a weak point of the current literature. A systematic study of

14-3-3 dimer formation requires an assay system that allows for medium to high

throughput testing of interactions to cope with the large number of 14-3-3

combinations. Furthermore such a system should allow for the identification of

subcellular localisations without bias, which can result when performing fractionated

protein extraction. Finally, to be biologically meaningful, this assay system should

allow for in planta analysis, ideally in the organism from which the genes of interest

where isolated. BiFC analysis as described in Chapter 3 and by Walter and

co-workers (Walter et al., 2004) has the potential to fulfil all of the above-summarised

requirements.

The work presented in this chapter addresses the questions discussed above

with regards to formation and localisation of Arabidopsis 14-3-3 dimers. The

transient BiFC assay system, developed as described in Chapter 3, was employed in


combination with the 14-3-3 BiFC vector library (Section 3.2.4) to investigate these

questions. In detail, the aims of this work were:

(1) To establish whether Arabidopsis 14-3-3 isoforms can freely form dimers with each

other or whether preferences exist for some isoforms to form homo- or

heterodimers as was found in animals.

(2) To identify the subcellular localisations of all 14-3-3 dimer combinations tested

(3) To determine potential dimer preferences with regards to subcellular structures

which can then serve as the basis for functional analysis.

4.1.3. Interaction of 14-3-3s with target proteins

In addition to the above, studies into the interaction of a subset of 14-3-3s with

a target protein were used to complement the dimerisation and localisation studies

(described in Section 4.2.6). The target protein was chosen based on the outcome of

dimerisation and localisation studies presented in this chapter (Sections 4.2.2 to 4.2.4).

The focus was on potential interactions of 14-3-3s with nuclear target proteins.

The literature reports several potential roles for 14-3-3 proteins in the nucleus,

from interaction with transcription factor complexes to shuttling of proteins between

the nucleus and the cytosol (reviewed in Eckardt, 2001; Comparot et al., 2003;

Roberts, 2003; Dougherty and Morrison, 2004; Chevalier et al., 2009). Plant 14-3-3

proteins were identified in protein complexes that bind to the G-box elements found

in promoters of diverse plant genes, such as the alcohol dehydrogenase gene (Lu et al.,

1992), suggesting functional associations of 14-3-3s with transcription factors. A

large number of transcription factors in plants have been shown to interact with

14-3-3 proteins, including EmBP1/VIVIPAROUS1 (Schultz et al., 1998), TBP2 (TATA box

binding protein; Pan et al., 1999), TFIIB (transcription factor IIB; Pan et al., 1999),

ZMHOX1a/ZMHOX1b (Zea mays homeobox; Halbach et al., 2000), RSG (Repression of


Shoot Growth; Igarashi et al., 2001), HvABI5 (Hordeum vulgare ABA insensitive 5;

Schoonheim et al., 2007) and BZR1 (brassinazole resistant-1; Gampala et al., 2007).

Thus, potential roles for 14-3-3s in nuclear functions are well established.

A wider literature search showed at least twenty proteins are either shown or

predicted to interact with 14-3-3 proteins in the nucleus (Chang et al., 2009; Paul et al.,

2009). The nuclear 14-3-3 interactors include members of a small, plant-specific

family of histone deacetylase proteins in Arabidopsis, the histone deacetylase 2 (HD2)

family (Paul et al., 2009). In mammalian cells, histone deacetylases are found to

interact with 14-3-3 proteins, and such interactions sequester HDACs in the cytoplasm

(Grozinger and Schreiber, 2000; Wang et al., 2000). Similar cytoplasmic

sequestration was shown when 14-3-3 proteins bound plant nuclear proteins, such as

RSG and BZR1 (Igarashi et al., 2001; Gampala et al., 2007). To date, it is unclear if

such cytoplasmic sequestration by 14-3-3s also occurs as part of the regulation of the

plant-specific HD2s in vivo.

The Arabidopsis genome encodes 18 HDAC genes which can be categorised into

three types of HDACs: RPD3-like (also known as HD1), HD-tuin (also known as HDT

and HD2) and sirtuin (Hollender and Liu, 2008). The HD2 subfamily comprises 4

members: HD2A, HD2B, HD2C and HD2D (also known as HDT1, HDT2, HDT3 and HDT4,

respectively; Pandey et al., 2002; Hollender and Liu, 2008). The HD2 family is a

plant-specific class of HDACs, which is not found in animals or fungi (Pandey et al.,

2002; Hollender and Liu, 2008). Transcription patterns of all four HD2s share similar

profiles; all are highly expressed in inflorescence tissues but are under-expressed in

vegetative tissues, pollens, seeds and late-stage flowers (Zhou et al., 2004; Hollender

and Liu, 2008). GFP fusion approaches indicate that HD2A, HD2B and HD2C all

localise exclusively to the nucleolus (Lawrence et al., 2004; Zhou et al., 2004; Pendle

et al., 2005; Sridha and Wu, 2006). Functional analyses revealed that HD2A is


involved in reproductive development. Antisense HD2A transgenic plants exhibit a

phenotype of stunted siliques (Wu et al., 2000), whilst overexpression of GFP-HD2A

resulted in abnormal leaf morphology, flowers with shorter filaments, aborted seed

development, sterility and late flowering (Lawrence et al., 2004). HD2A and HD2B

act together in leaf polarity determination, possibly by controlling the levels or

distribution of two micro RNAs involved in abaxial/adaxial axis formation (Ueno et al.,

2007). In contrast, little is known about the biological function of HD2C and HD2D.

It is reported that HD2C is involved in abscisic acid and abiotic stress responses,

including drought and salinity (Sridha and Wu, 2006). Furthermore, HD2A, HD2B

and HD2C were identified in 14-3-3 protein complexes using an interactomic assay

(Paul et al., 2009). This report provided, for the first time, evidence for an

interaction of 14-3-3 proteins with HDACs, indicating that these plant-specific HDACs,

similar to their mammalian relatives, may be regulated by 14-3-3s. However, the

roles of 14-3-3 in regulating HD2s are still elusive. Recently, our lab was able to

demonstrate interaction of two 14-3-3 isoforms (chi and kappa) with HD2C using a

transient BIFC approach (M. van der Kwast, unpublished PhD thesis). Interaction was

observed in the nucleus and nucleolus but was absent from the cytoplasm.

Here HD2C (At5g03740) was used to test for interaction with a selection of

14-3-3 proteins to elucidate whether:

(1) 14-3-3s, other than the previously tested chi and kappa isoforms, are able to

interact with this HDAC, i.e. is there isoform specificity;

(2) the localisation preference of a 14-3-3 protein can impact on its ability to interact

with HD2C or if

(3) the preference of a 14-3-3 protein for nuclear or cytosolic localisation is

determined by the target protein, in this case HD2C.


4.2. Results

4.2.1. A database analysis of the subcellular localisation of Arabidopsis

14-3-3 proteins using the SUBcellular location database for

Arabidopsis proteins (SUBA)

A database analysis based on SUBA data was performed to establish a current

view of the subcellular localisations of Arabidopsis 14-3-3 isoforms before

commencing with 14-3-3 dimerisation assays using BiFC (Table 4-1). Such

localisation data could provide essential background knowledge for an informed

interpretation of 14-3-3 BiFC assays. The SUBA localisation data are based on in

silico predictions of protein localisation as well as on experimental approaches such as

proteomic studies and GFP-fusion approaches (Heazlewood et al., 2007).

Data retrived from the SUBA database indicated that all Arabidopsis 14-3-3

members localise to the cytosol by LOCtree or SubLoc predictions (Table 4-1).

Similarly, in addition to cytosolic localisation, eight 14-3-3 isoforms, chi, iota, kappa,

lambda, nu, pi, psi and upsilon, may also localise to the nucleus, based on the

predictors AmiGO or WoLF PSORT (Table 4-1). The same programs also predicted

that only 14-3-3 chi, epsilon and omega localise to the plasma membrane whilst

14-3-3 epsilon, mu and phi are predicted to localise to plastids as well. Two isoforms,

14-3-3 lambda and omicron, were predicted to be found in peroxisomes by PeroxiP

and WoLF PSORT, respectively (Table 4-1).

The SUBA data also indicated that proteomic data sets were available for ten of

the Arabidopsis 14-3-3 proteins whilst subcellular localisation data as identified by

fluorescent fusion protein approaches were available for six isoforms (Table 4-1).

None of the experimental datasets covered the localisation of the three 14-3-3


Table 4-1. Subcellular localisation of Arabidopsis 14-3-3 proteins using the

SubCellular Proteomic Database [SUBA; Heazlewood et al. (2007)].

The table summarises the search outcomes for the subcellular localisation of the Arabidopsis

14-3-3 protein family using predictor program and experimental datasets based on

proteomics and fluorescent protein experiments as available in the SUBA database.

14-3-3 isoform

Localisation Predicted by

program

References for proteomic

localisation data

References for fluorescent protein

localisation data

chi

cytosol nucleus plasma membrane plastid

[1], [2] [3] [4] -

- [10] [7], [8], [11], [15]

- - - -

epsilon cytosol plasma membrane plastid

[1], [2] [3] [4]

- [8] -

- - -

iota cytosol nucleus

[1], [2] [4]

- -

- -

kappa cytosol nucleus plastid

[2] [3] -

- [10] [11]

- [9] -

lambda

cytosol nucleus plasma membrane plastid peroxisome

[1], [2] [3], [4] - - [5]

- - [8] [11] [16]

[12] [9], [12] - - -

mu

cytosol nucleus plasma membrane plastid peroxisome

[1], [2] - - [3] -

- - [7] - [16]

[6], [16] [6] [16] - -

nu

cytosol nucleus plasma membrane plastid

[1], [2] [3], [4] - -

- - [7], [8], [13] [11]

- - - -

omega

cytosol nucleus plasma membrane cytoskeleton vacuole peroxisome

[1], [2] - [4] - - -

- - [8] - [14] [16]

[9] [9] - [9] - -

omicron cytosol peroxisome

[1], [2] [4]

- -

- -

phi

cytosol nucleus plasma membrane cytoskeleton plastid

[1], [2] - - - [4]

- - [8] - -

[9] [9] - [9] -

pi cytosol nucleus

[1], [2] [4]

- -

- -

psi

cytosol nucleus plasma membrane peroxisome

[1], [2] [4] - -

- - [7], [8] [16]

[16] - [16] -

upsilon cytosol nucleus plasma membrane

[1], [2] [3], [4] -

- - [7], [8]

- - -

“-“: No hit from the database search. References: [1] LOCtree [http://www.rostlab.org/services/LOCtree/; Nair and Rost (2005)]; [2] SubLoc [http://www.bioinfo.tsinghua.edu.cn/SubLoc/; Hua and Sun (2001)]; [3] AmiGO [http://amigo.geneontology.org/; Carbon et al. (2009)]; [4] WoLF PSORT [http://wolfpsort.org/; Horton et al. (2007)]; [5] PeroxiP [http://www.bioinfo.se/PeroxiP/; Emanuelsson et al. (2003)]; [6] Koroleva et al. (2005); [7] Marmagne et al. (2007); [8] Benschop et al. (2007); [9] Paul et al. (2005); [10] Bae et al. (2003); [11] Kleffmann et al. (2004); [12] Rienties et al. (2005); [13] Karlova et al. (2006); [14] Jaquinod et al. (2007); [15] Zybailov et al. (2008); [16] Reumann et al. (2009). Date of the database search: 13/5/2009.

http://www.rostlab.org/services/LOCtree/

http://www.bioinfo.tsinghua.edu.cn/SubLoc/

http://amigo.geneontology.org/

http://wolfpsort.org/

http://www.bioinfo.se/PeroxiP/


isoforms, iota, omicron and pi. Experimental datasets based on proteomic studies

and GFP-fusion approaches are only partially in agreement with the predicted

subcellular localisations of 14-3-3 isoforms. For example, of the eight 14-3-3

isoforms predicted to localise to the nucleus (Table 4-1), only 14-3-3 chi and kappa

were experimentally identified in the nuclear proteome (Bae et al., 2003). Similarly,

although only two isoforms, lambda and omicron, were predicted to localise to

peroxisomes (Table 4-1), Reumann and co-workers demonstrated using a proteomics

approach that 14-3-3 lambda, mu, omega and psi localise to this organelle (Reumann

et al., 2009). Furthermore, 14-3-3 chi, kappa, lambda and nu were found in the

chloroplast proteome (Kleffmann et al., 2004; Zybailov et al., 2008) but none of these

four isoforms were predicted to localise to plastids (Table 4-1).

14-3-3 GFP fusion approaches are only available for six Arabidopsis 14-3-3

isoforms, kappa, lambda, mu, omega, phi and psi (Koroleva et al., 2005; Paul et al.,

2005; Rienties et al., 2005; Reumann et al., 2009; Table 4-1). These revealed that

even very similar 14-3-3 isoforms such as 14-3-3 kappa and lambda, which are 92%

identical at the amino acid level (Wu et al., 1997a), show differences in subcellular

localisation. In transgenic Arabidopsis plants, the 14-3-3 kappa-GFP fusion protein

was found predominantly in nuclei of trichomes and guard cells, whilst the

lambda-GFP protein localised to nuclei of trichomes but not to nuclei of guard cells

(Paul et al., 2005). In addition, the omega-GFP and phi-GFP fusion proteins were

evenly distributed between the cytosol and nuclei in the cells of trichomes and guard

cells (Paul et al., 2005).

A dependence of 14-3-3 localisation based on the experimental system is also

found for some 14-3-3s. For example, YFP-14-3-3 lambda fusion proteins were

found in the nuclei of transiently transformed cowpea protoplasts (Rienties et al.,

2005), which was consistent with the localisation of lambda-GFP observed in


Arabidopsis trichomes (Paul et al., 2005). However, in contrast to its location in

Arabidopsis trichomes (Paul et al., 2005), lambda also localised to the cytosol of

cowpea protoplasts (Rienties et al., 2005), indicating a clear dependence on the

experimental system and/or the cell type investigated. A dependence on the

experimental system became more obvious when data for 14-3-3 mu were analysed.

In transiently transformed Arabidopsis suspension cells, GFP-mu fusion proteins

localised to nuclei and the cytosol (Koroleva et al., 2005). In contrast, Reumann and

co-workers found 14-3-3 mu-YFP fusion proteins localised to the cytosol and/or the

plasma membrane, but it was absent from the nucleus of transiently transformed

tobacco epidermal cells (Reumann et al., 2009).

It can be concluded that the cytosol is a common location for 14-3-3 isoforms.

Thus, it can be expected that BiFC assays should reveal 14-3-3 dimerisation via

YFP-based fluorescence at least in the cytosol. Predicted and experimental datasets

indirectly suggest that certain 14-3-3 dimer combinations are found in some

organelles. As discussed above, a number of 14-3-3s localise to the nucleus as GFP

fusion proteins. However, several 14-3-3 isoforms (epsilon, iota, lambda, mu, nu,

omega, omicron, phi, pi, psi and upsilon) were not identified in the nucleus when

using proteomics approaches (Table 4-1). Of these, epsilon, mu, omega, omicron

and phi were not predicted to localise to the nucleus. Thus, according to the SUBA

dataset, it is unlikely that these isoforms will be found as part of a 14-3-3 dimer within

the nucleus. The BiFC analysis described in Chapter 3 will interrogate this and be

used to verify or to refute the information in available datasets. Furthermore, none

of the datasets predicted or showed localisation of 14-3-3 poteins in Golgi, ER or the

nucleoli. Therefore, the BiFC experiments conducted during this thesis paid special

attention to those organelles and subcellular localisations.


4.2.2. Arabidopsis 14-3-3 isoforms can form dimers with each other in

living plant cells

To perform the systematic survey of dimerisations among the ten Arabidopsis

14-3-3 isoforms, each of the possible fifty-five 14-3-3 dimer combinations was tested

individually by co-infiltrating a suspension mixture of Agrobacteria cells into Nicotiana

leaves. The Agrobacterium mixture contained three strains: a 14-3-3-YN strain, a

14-3-3-YC strain and a silencing suppressor-p19 carrying strain (Materials and

Methods, Section 2.4.5). For example, the combination of chi-YN and chi-YC strains

for co-infiltration was used to examine 14-3-3 chi homodimerisation (chi-chi); while

the combination of chi-YN and epsilon-YC was used to establish heterodimerisation

between the 14-3-3 chi and epsilon isoforms (chi-epsilon). BiFC fluorescence, due to

re-constitution of YFP upon 14-3-3 dimerisation, was detected for all examined 55

possible dimer combinations. The results indicated that all ten Arabidopsis 14-3-3

isoforms were able to dimerise with each other in vivo (Fig. 4-1). A representative

micrograph of the BiFC fluorescence for each 14-3-3 dimeric combination is shown in

Fig. 4-1. There was no obvious preferential dimerisation among the ten Arabidopsis

14-3-3 isoforms from the BiFC data.

The BiFC fluorescence not only demonstrated 14-3-3 dimerisations but also

indicated the subcellular locations of the dimers. Generally, the BiFC fluorescence

was observed in the cytosol and the nucleus. Further analyses of the 14-3-3 dimer

subcellular localisations are described later (Section 4.2.3).

The effect of YN and YC fusion configurations on 14-3-3 dimerisations and

localisation was also investigated. For example, did a dimer composed of

chi-YN/kappa-YC result in the same outcome as a dimer composed of kappa-YN/chi-YC.

In addition, no obvious differences for the two possible YN/YC configurations

chi epsilon iota kappa mu nu omega omicron phi psi

psi

phi

omicron

omega

nu

mu

kappa

iota

epsilon

chi

N

N

N

N

N

N

N

N

N

N

N

N

N

NN

N

N

N

N NN

N

N

N

N

N

N

N

N

NN

N N

NN

N

N

N

N

N

N

N

N

N

N

N

N

N

N

N

N

N

N

N

N

Fig. 4-1. Dimerisation of Arabidopsis 14-3-3 proteins using transient BiFC assays in N.benthamiana leaves.

Ten Arabidopsis 14-3-3 isoforms were tested for their ability to form homo- and heterodimersemploying transient BiFC assays in N. benthamiana leaf epidermal cells. The ability of each 14-3-3 protein to homodimerise and to form dimers with all other isoforms is shown by green YFPfluorescence. Dimers were identified in the nucleus and cytoplasm. The micrographs representone of several possible subcellular localisation patterns observed for individual 14-3-3 dimers(see also Section 4.2.3). The total number of cells examined for individual 14-3-3 dimer aresummarised in Table 4-2. N, nucleus. Scale bars = 50 μm.



were observed for the dimer combinations chi/omega, chi/mu, mu/epsilon, mu/iota

and mu/kappa (data not shown). Thus, it was assumed that dimers of the form

A-YN/B-YC and B-YN/A-YC were equivalent and the analysis was not extended to all

possible 14-3-3 dimer combinations.

4.2.3. Subcellular localisation of 14-3-3-BiFC dimeric complexes

4.2.3.1. Dual cytosolic/nuclear localisation of 14-3-3 dimers

The subcellular localisation for each 14-3-3 dimeric combination was

determined from the fluorescence micrographs. Fluorescence was detected in the

cytosol and in cytoplasmic strands within cells, indicating that all the dimers are

cytosolic under the experimental conditions used in this study (Fig. 4-1). Additionally,

BiFC fluorescence was detected in the nucleus of most of the cells showing

fluorescence (Fig. 4-1), indicating nuclear localisation of 14-3-3 dimers. All fifty-five

14-3-3 dimers showed a similar pattern of dual cytosolic and nuclear localisation.

However, the intensity of the fluorescence in the nucleus varied with the dimer

combination and sometimes also with the cell. For example, strong nuclear

fluorescence was often observed in cells expressing mu-psi heterodimers or kappa

homodimers (Fig. 4-1). In contrast, nuclear fluorescence was generally weak in cells

expressing psi-omega heterodimers or nu homodimers (Fig. 4-1).

A small structure (sometimes more than one), which showed no fluorescence

and was possibly the nucleolus, was found in the nucleus in all the cells analysed

across the 55 dimeric combinations tested. This suggests that 14-3-3 dimers

localised to the nucleus are absent from the nucleolus (Figs. 4-1, 3-12 B, 3-13 and data

not shown).

In some cases, fluorescence was also observed in a thin layer surrounding the

nucleus, as shown in the micrographs of epsilon-nu, nu-nu and psi-nu dimers (Fig. 4-1)


as well as an omega-nu dimer shown in Fig. 4-2 B, suggesting that some 14-3-3 dimers

localise to the nuclear envelope/ER.

The different localisation patterns observed for 14-3-3 dimers were used to

establish a classification of 14-3-3 dimer localisation (Section 4.2.3.3). Special

emphasis was placed on the intensity of the observed fluorescence in the nuclei.

This classification was then used to interrogate a larger number of cells for each of the

individual dimer combinations to investigate whether the observed patterns were

consistent and dimer specific (Section 4.2.4).

4.2.3.2. 14-3-3 dimers were absent from chloroplast stroma but may be

associated with chloroplastic membranes

Chloroplastic localisation was predicted for the epsilon, mu and psi isoforms and

a proteomics approach localised the 14-3-3s chi, kappa, lambda and nu to plastids

(see Section 4.2.1 for details and references). Additionally, 14-3-3 isoforms epsilon,

mu, nu and upsilon have been shown to localise to chloroplasts (Sehnke et al., 2000).

However, none of the 14-3-3 isoforms mentioned above contains a transit peptide.

In the transient BiFC assay in Nicotiana leaf epidermal cells, the nucleus was

usually surrounded by some chloroplasts (Figs. 4-1, 3-12 and 3-13). Thus, this assay

system was used to investigate if 14-3-3 isoforms (especially the predicted or reported

isoforms to be chloroplastic) can dimerise within the chloroplasts in the epidermal

cells.

As described in Section 3.2.7, BiFC fluorescence was not detectable within

chloroplasts for the 14-3-3 kappa homodimer (Figs. 3-12 C and 3-13 A-D) and

phi-omega heterodimer (Fig. 3-13 E-H). When the dimerisation assay was expanded

to other 14-3-3 isoforms, including those previously predicted/reported to be

chloroplastic (chi, epsilon, kappa, mu, nu and psi), none of the possible dimer

combinations demonstrated any visible BiFC fluorescence within chloroplasts (Fig. 4-1


and data not shown). Instead, BiFC fluorescence was clearly visible around

chloroplasts for a large number of dimer combinations (Figs. 4-1, 4-2 C, 3-12 and 3-13).

This may indicate an association of 14-3-3 dimers with the outside of chloroplasts or

localisation within the chloroplastic membranes or the chloroplastic intermembrane

space.

4.2.3.3. Variable nuclear localisation patterns of 14-3-3 dimers were

observed

Variable fluorescence intensities in the nucleus were observed not only across

different 14-3-3 dimeric combinations (Fig. 4-1) but also between different cells

expressing the same dimeric combination (Fig. 4-2). The differences were used to

classify nuclear localisation patterns of 14-3-3 dimers and to investigate whether the

observed patterns were dimer specific. To this end, the relative levels of nuclear to

cytoplasmic fluorescent signals were quantified for a large number of cells from

micrographs of each 14-3-3 dimerisation experiment using ImageJ software (Section

2.5.3.1). The ratios of nuclear to cytoplasmic BiFC fluorescence intensity (N/C) from

individual cells were used to define three nuclear localisation patterns. These were

as follows:

Nuclear Pattern 1: Cells were classified as showing Pattern 1 if the N/C ratio was

greater than 1.5, indicating prominently nuclear localisation of 14-3-3 dimers. This

matched the optical observation.


in between 0.7 to 1.5, indicating relative equal distribution of fluorescence between

the cytosol and the nucleus. Again, optical observations were used to support this.

Fig. 4-2. Variable distribution patterns of BiFC fluorescence by 14-3-3 dimerisation observedin transiently transformed Nicotiana epidermal cells.

Variable distribution patterns of BiFC fluorescence by 14-3-3 dimerisation were observedbetween and within dimer combinations as demonstrated here for the heterodimerisation of14-3-3 omega-YN and nu-YC fusion proteins in Nicotiana epidermal cells. Nuclear versuscytosolic fluorescence ratios (N/C ratios), obtained with image analysis, were used to classifythese patterns. A. Some cells had stronger fluorescence in the nucleus than in the cytosol (N/C> 1.5: Pattern 1). B. Other cells had apparently equal levels of fluorescence in the nucleus andcytosol (0.7 ≤ N/C ≤ 1.5: Pattern 2). C. A third category of cells showed stronger fluorescence inthe cytosol than in the nucleus (N/C < 0.7: Pattern 3). Bright field images of A, B and C areshown in D, E and F, respectively. Cyt, cytosol; Chl, chloroplast; N, nucleus; NE, nuclearenvelope. Scale bars = 50 μm.

A D

B

Pattern 2

Pattern 3

Pattern 1

C F

E

N

N

N

N/C=0.8

N/C=0.4

N/C=2.3


Cyt

Cyt

Cyt

NE

Chl

Chl




less than 0.7, representing weak fluorescence to almost absence of fluorescence from

the nucleus.

The classification scheme was applied to all cells photographed for each of the

fifty-five dimer combinations (Table 4-2). Using the heterodimerisation of 14-3-3

omega-nu (omega-YN/nu-YC) as an example, for which 25 cells were analysed, three

cells belonged to the Pattern 1, twelve cells were in the Pattern 2 category and ten

cells appeared in the Pattern 3 category (Table 4-2). Representative micrographs for

each of the patterns observed for the omega-nu dimer are shown in Fig. 4-2.

To provide an overall view of the nuclear localisation patterns for each 14-3-3

dimer, a scatterplot analysis was performed (Fig. 4-3). In total 581 cells (across 55

dimerisations) were analysed. Of these, 182 cells (31 %) were in the Pattern 1

category, 347 cells (60 %) in the Pattern 2 and 52 cells (9 %) were in the Pattern 3

category. Although the frequency of the three categories is dependent somewhat

on the number of cells analysed for each dimer, as some dimers appeared

preferentially in one over the other categories, the overall data indicated that the

most prominent pattern was Pattern 2, with almost twice as many cells found

exhibiting this pattern than in the next frequent category, Pattern 1, and over six times

more frequent than Pattern 3.

4.2.4. Some 14-3-3 dimers showed preferential subcellular localisations

The number of cells falling into one of the three N/C categories is summarised

for each of the fifty-five dimers in Table 4-2. Although for most dimers the number

of cells used to do the quantification analysis was far too low to be of statistical

significance, a trend suggesting preferential localisation patterns was observed for a

number of 14-3-3 dimers (Figs. 4-4 to 4-5).


Table 4-2. Quantitative analysis of the subcellular localisation of Arabidopsis 14-3-3

dimers as determined by image analysis of transient BiFC assays.

Nuclear localisation patterns were determined from epifluorescence images of all

possible dimer permutations of ten Arabidopsis 14-3-3 isoforms. The ratios of

nuclear to cytoplasmic (N/C) BiFC fluorescence intensities were estimated and used to

classify the cells into Pattern 1 (N/C ratio > 1.5), indicating prominently nuclear

localisation of 14-3-3 dimers, Pattern 2 (N/C ratio 0.7 to 1.5), indicating near equal

distribution between nucleus and cytosol and nuclear Pattern 3 (N/C ratio < 0.7),

representing weak or lack of nuclear fluorescence. The number of cells exhibiting

the three patterns (1, 2, and 3) and the total number of cells analysed (total) is given

for each 14-3-3 dimer. Homodimer combinations of 14-3-3s are highlighted in grey

for ease of navigation.

14-3-3 dimer

Localisation pattern

14-3-3 dimer

Localisation pattern

1 2 3 total 1 2 3 total

chi-chi 3 12 - 15 omicron-chi 5 5 - 10

chi-epsilon - 7 - 7 omicron-epsilon - 7 - 7

chi-kappa 5 8 - 13 omicron-iota 6 3 - 9

chi-nu - 5 3 8 omicron-kappa 5 2 - 7

chi-omega - 7 - 7 omicron-mu 4 4 - 8

epsilon-epsilon 3 6 - 9 omicron-nu - 12 - 12

epsilon-kappa 6 - - 6 omicron-omega 1 5 1 7

epsilon-nu 1 19 - 20 omicron-omicron 2 4 - 6

iota-chi - 3 - 3 phi-chi 1 6 - 7

iota-epsilon 1 9 - 10 phi-epsilon 3 5 - 8

iota-iota 1 11 - 12 phi-iota 1 11 - 12

iota-kappa - 6 - 6 phi-kappa 7 1 - 8

iota-nu 2 2 7 11 phi-mu 3 2 - 5

kappa-kappa 30 1 - 31 phi-nu - 9 2 11

kappa-nu 1 10 - 11 phi-omega - 6 - 6

mu-chi 3 7 - 10 phi-omicron 3 4 - 7

mu-epsilon 15 20 - 35 phi-phi 4 17 3 24

mu-iota 4 5 - 9 psi-chi 3 4 - 7

mu-kappa 14 - - 14 psi-epsilon 2 2 1 5

mu-mu 4 10 - 14 psi-iota 3 1 1 5

mu-nu - 6 - 6 psi-kappa 4 - - 4

nu-nu 2 12 19 33 psi-mu 8 1 - 9

omega-epsilon 2 3 - 5 psi-nu - 2 3 5

omega-iota - 5 - 5 psi-omega - 2 2 4

omega-kappa 2 4 6 psi-omicron 7 7 - 14

omega-mu 3 18 - 21 psi-phi 5 7 - 12

omega-nu 3 12 10 25 psi-psi - 4 - 4

omega-omega - 6 - 6

1: chi-chi 2: chi-epsilon 3: chi-kappa 4: chi-nu 5: chi-omega

6: epsilon-epsilon 7: epsilon-kappa 8: epsilon-nu 9: iota-chi 10: iota-epsilon

11: iota-iota 12: iota-kappa 13: iota-nu 14: kappa-kappa 15: kappa-nu

16: mu-chi 17: mu-epsilon 18: mu-iota 19: mu-kappa 20: mu-mu

21: mu-nu 22: nu-nu 23: omega-epsilon 24: omega-iota 25: omega-kappa

26: omega-mu 27: omega-nu 28: omega-omega 29: omicron-chi 30: omicron-epsilon

31: omicron-iota 32: omicron-kappa 33: omicron-mu 34: omicron-nu 35: omicron-omega

36: omicron-omicron 37: phi-chi 38: phi-epsilon 39: phi-iota 40: phi-kappa

41: phi-mu 42: phi-nu 43: phi-omega 44: phi-omicron 45: phi-phi

46: psi-chi 47: psi-epsilon 48: psi-iota 49: psi-kappa 50: psi-mu

51: psi-nu 52: psi-omega 53: psi-omicron 54: psi-phi 55:psi-psi

Fig. 4-3. Scatter plot analysis of localisation patterns of 14-3-3 dimers as determined bytransient BiFC assays and image analysis software.

Micrographs of cells expressing two 14-3-3 BiFC constructs were chosen for analysis offluorescence intensity using imageJ software (Section 2.5.3.1). The ratios of nuclear tocytosolic BiFC fluorescence intensity (N/C) obtained for individual cells were plotted for thecorresponding 14-3-3 dimer combinations. Each point, represented in the graph by diamonds,shows the N/C fluorescent ratio for a single cell. Values of the ratios above 1.5 represent cellsthat had a stronger fluorescence in the nucleus than in the cytosol (Pattern 1). The N/C ratiosbetween 1.5 and 0.7 correspond to cells in which the fluorescence intensity in the cytosol andnuclei appeared similar (Pattern 2). Values below 0.7 represented cells that had strongerfluorescence in the cytosol than in the nucleus (Pattern 3; see text for detail).

14-3-3 dimer combinations

Nu

cle

ar/

cyto

pla

sm

icsig

nal ra

tio

Pattern 1

Pattern 2

Pattern 3



To simplify the analysis, the fluorescence values were plotted to allow either

comparison of homodimers or of dimers with one common 14-3-3 protein (Figs. 4-4

and 4-5). When comparing the values of ten 14-3-3 homodimers, it appeared that

cells containing the dimers epsilon-epsilon, mu-mu, omicron-omicron, iota-iota,

chi-chi and kappa-kappa exhibited both Patterns 1 and 2; cells expressing the dimers

omega-omega and psi-psi exhibited only the Pattern 2 category; and cells containing

the dimers nu-nu and phi-phi displayed all three fluorescence patterns (Fig. 4-4).

Apparently, cells expressing the kappa homodimer exhibited a strong preference for

Pattern 1, with 30 of the 31 analysed cells having an N/C fluorescent ratio of more

than 1.5 (Fig. 4-4 and Table 4-2). In strong contrast, few cells containing the nu

homodimer showed Pattern 1 fluorescence, with less than 10% of the cells (2 out of

33) having an N/C fluorescence ratio of more than 1.5, and most of the cells having an

N/C ratio of less than 1 (Fig. 4-4 and Table 4-2). The majority of cells containing the

remaining 14-3-3 homodimers exhibited Pattern 2 (Fig. 4-4 and Table 4-2).

As for cells containing the 14-3-3 heterodimers, some exhibited the Pattern 1

only, such as those containing mu-kappa (14 cells out of 14) and epsilon-kappa (6 cells

out of 6), and others demonstrated Pattern 2, such as those expressing chi-epsilon (7

cells out of 7) and iota-kappa (6 cells out of 6; Table 4-2). Besides these, cells

expressing other heterodimer combinations displayed multiple patterns such as cells

expressing mu-epsilon, which showed both Patterns 1 and 2, with almost equal

distribution between the two (15 cells in Pattern 1 and 20 cells in Pattern 2; Table

4-2).

Only a few dimer combinations resulted in cells exhibiting Pattern 3 labelling

(weak labelling of the nucleus). None of these were solely found with Pattern 3

labelling (Table 4-2). For example, cells expressing nu homodimers demonstrated all

three patterns although with a bias towards Patterns 2 and 3 (Pattern 1: two cells,


Fig. 4-4. Subcellular localisation of 14-3-3 homodimers as determined by transient BiFC

assays and image analysis software.

A subset of the nuclear to cytosolic fluorescence ratios shown in Figure 4-3, focusing on

14-3-3 homodimers, are shown to demonstrate trends in the subcellular distributions of these

dimers. For details see Figure 4-3.

Pattern 2: twelve cells and Pattern 3: nineteen cells). Similarly, cells with the

omega-nu heterodimer exhibited all three patterns with an underrepresentation of

Pattern 1 (Pattern 1: three cells, Pattern 2: twelve cells and Pattern 3: ten cells, Table

4-2). Interestingly, six out of the eleven dimers containing the 14-3-3 nu isoform

resulted in cells displaying Pattern 3 (Table 4-2).

A trend towards preferential localisation patterns for the kappa- and nu-involved

14-3-3 dimers was apparent (Fig. 4-5). Of the ten kappa-involved dimers, cells

expressing six of them had a high propensity to be in Pattern 1; while those of the

remaining four (chi-kappa, iota-kappa, kappa-nu and omega-kappa) had preference to

be in Pattern 2 (Fig. 4-5). None of the cells with the ten kappa-involved dimers were

observed in Pattern 3 (Fig. 4-5 and Table 4-2). By contrast, cells expressing eight out

of the ten nu-related dimers were mainly in Pattern 2 and those with the remaining


Fig. 4-5. The impact of 14-3-3 isoforms kappa and nu on the subcellular localisation of

14-3-3 dimers.

A subset of the nuclear to cytosolic fluorescence ratios shown in Figure 4-3, focusing on

14-3-3 dimers containing 14-3-3 kappa or 14-3-3 nu, were shown to demonstrate the impact

of these 14-3-3 isoforms on the localisation of their homo- and heterodimers. For details

see Figure 4-3.

two dimers, iota-nu and nu-nu, were observed a high propensity to be in Pattern 3 (fig.

4-5 and Table 4-2). Although, these trends are somewhat influenced by the number

of cells surveyed for individual dimers, an overall tendency is apparent, whilst the

involvement of 14-3-3 kappa strongly promotes nuclear localisation of 14-3-3 dimers,

14-3-3 nu obstructs or reduces it (Fig. 4-5).

4.2.5. Discovery of a novel localisation pattern of 14-3-3 dimers

When analysing a larger number of cells for localisation patterns, a novel

distribution of 14-3-3 dimers was discovered. Fluorescence was observed in distinct,

punctuated areas within the cytoplasm, as shown in the selected images in Fig. 4-6.

This distinct 14-3-3 dimer labelling was different from the usually diffuse fluorescence

Fig. 4-6. Epifluorescence images demonstrating a novel, distinct cytoplasmic localisationpattern of 14-3-3 dimers in transiently transformed Nicotiana benthamiana leafepidermal cells.

Some Nicotiana epidermal leaf cells transiently expressing the indicated 14-3-3 BiFCconstructs showed a distinct cytoplasmic distribution pattern. This pattern was manifestedas punctuate, fluorescent bodies, which were observed at low frequency in transformedcells. This pattern was designated as Pattern 4 (see text for detail). Epifluorescence imageswere obtained using a fluorescence microscope equipped with a GFP filter, except for theiota-nu image (F) which was photographed using a FITC filter instead of the GFP filter. N,nucleus. Scale bars = 10 μm.

iota-chi chi-omega mu-epsilon

iota-nu omicron-iota omicron-omega

phi-nu omega-omega phi-phi

A B C

D E F

G H I

N

N

N

N



observed in the cytoplasm of most cells showing BiFC fluorescence (Fig. 4-1).

Although this novel pattern was observed for many 14-3-3 dimers, it was usually

underrepresented when compared to the frequency of the diffuse cytoplasmic

distribution. The approximate frequency of cells showing this novel pattern was

between 1% and 3.3% (data not shown).

The novel distribution pattern always occurred concurrently in cells showing

either nuclear Pattern 1 or 2, but never in cells with the Pattern 3 (data not shown).

For identification purposes, the novel cytoplasmic pattern is designated as Pattern 4

(Table 4-3). This pattern will be further analyzed in chapter 6.

The localisation patterns for all 14-3-3 dimeric combinations are summarised in

Table 4-3. From this it is apparent that the great majority of 14-3-3 dimers were

responsible for more than one localisation pattern. The novel Pattern 4 was

observed in cells containing 27 of the 55 analysed dimeric combinations, suggesting

the novel cytoplasmic localisation was possibly a general characteristic of 14-3-3

dimers and was not dimer-specific.

4.2.6. Investigation into the interaction of 14-3-3 proteins with the histone

deacetylase HD2C

The coding region of HD2C was cloned by Michael van der Kwast (unpublished

PhD thesis, Dr Martin’s laboratory, UWA) into the BiFC vectors described in Chapter 3.

The HD2C-YC construct was used to demonstrate interaction with 14-3-3 chi-YN and

kappa-YN in the nucleus and nucleolus using transient BiFC assays (Michael van der

Kwast, unpublished data).

This interaction study was extended to other 14-3-3 isoforms to test for

potential specificity of 14-3-3 isoforms towards the HD2C protein. The potential for

the preferential localisation of 14-3-3s to impact on their ability to interact with HD2C,


Table 4-3. Summary of subcellular distribution patterns of Arabidopsis

14-3-3 dimer combinations.

Arabidopsis 14-3-3 dimers were classified according to their distribution between the nucleus

and cytosol as described (Table 4-2) using nuclear to cytosolic (N/C) BiFC fluorescence

intensity ratios, which were correlated to observations made using a fluorescent microscope.

Nuclear Pattern 1 corresponded to prominently nuclear localisation, Pattern 2 corresponded

to equal distribution between nucleus and cytosol and Pattern 3 denotes mostly cytosolic

localisation. The table also shows 14-3-3 dimers appearing in a novel pattern (Pattern 4),

characterised by BiFC fluorescence appearing in distinct, punctuate fluorescent bodies

distributed throughout the cytosol. The Pattern 4, occurred with a very low frequency and

was only observed in combination with Patterns 1 and 2, never with the Pattern 3. With few

exceptions, 14-3-3 dimers appeared in more than one N/C pattern. Figures a to d show

typical epifluorescence images for each of the patterns and their corresponding N/C ratios.

Scale bars = 10 μm.

chi epsilon iota kappa mu nu omega omicron phi psi 14-3-3

1, 2 2 2, 4 1, 2 1, 2 2, 3 2, 4 1, 2 1, 2, 4 1, 2, 4 chi

1, 2 1, 2 4 1 1, 2, 4 1, 2, 4 1, 2 2 1, 2, 4 1, 2, 3 epsilon

1, 2 2 1, 2 1, 2, 3, 4 2 1, 2, 4 1, 2, 4 1, 2, 3, 4 iota

1, 2 1 1, 2, 4 1, 2 1, 2 1, 2 1, 4 kappa

1, 2, 4 2 1, 2, 4 1, 2 1, 2, 4 1, 2, 4 mu

1, 2, 3, 4 1, 2, 3, 4 2, 4 2, 3, 4 2, 3 nu

2, 4 1, 2, 3, 4 2 2, 3 omega

1, 2 1, 2 1, 2, 4 omicron

1, 2, 3, 4 1, 2, 4 phi

2 psi

or whether HD2C determines the localisation of 14-3-3 proteins by recruiting them

into the nucleus or nucleolus were also addressed.

Experiments of 14-3-3-HD2C interactions were performed as described for the

14-3-3 dimerisation studies by co-agroinfiltration of Nicotiana leaves using the 14-3-3

isoforms chi, epsilon, kappa, mu, nu, omega and omicron as YN constructs paired with


HD2C-YC (Sections 2.4.4 and 4.2.2). The kappa and nu isoforms were chosen, as they

appeared to represent the two extremes with respect to strong or weak nuclear

localisation when analysed as 14-3-3 dimers (Section 4.2.4; Figs. 4-4 and 4-5). In

parallel, 14-3-3 homodimers were used as controls to enable direct comparison of the

localisation patterns between the 14-3-3 dimers and the 14-3-3-HD2C complexes.

All seven 14-3-3 isoforms tested were able to interact with HD2C (Fig. 4-7 and

data not shown). Additionally all the 14-3-3 HD2C interactions were found in the

nuclei and the nucleoli but were absent from the cytosol and other cellular organelles.

The interactions observed were independent of the nuclear localisation pattern of the

tested 14-3-3 homodimers. For example, as shown before (Figs. 3-12 C and 4-4), the

kappa homodimer showed dual localisation with stronger fluorescence in the nucleus

than in the cytosol (Pattern 1) and no fluorescence in the nucleolus (Fig. 4-7 B).

However, the 14-3-3 kappa-HD2C complex resulted in fluorescence in the nuclei and

nucleoli (Fig. 4-7 A). The nu homodimer, as shown before (Figs. 4-1 and 4-4), mainly

localised to the cytosol and was with only weak fluorescence in the nucleus (Fig. 4-7

D). In contrast, the 14-3-3 nu-HD2C complex was absent from cytoplasm and

exhibited strong fluorescence in nuclei and nucleoli (Fig. 4-7 C). The signals obtained

for interaction in the nucleolus were much stronger than those observed in the

nucleus. The remaining five 14-3-3 isoforms tested (chi, epsilon, mu, omega and

omicron), which as homodimers showed dual localisation in nucleus and cytosol (Figs.

4-1 and 4-4, Table 4-2 and data not shown), localised exclusively to the nuclei and

nucleoli when interacting with HD2C as described for the interactions of HD2C with

kappa and nu (Fig. 4-7 and data not shown).

YNYC

chi epsilon kappa mu nu omega omicron

HD2C + + + + + + +

Fig. 4-7. Interactions of 14-3-3 isoforms with histone deacetylase 2C (HD2C).

Protein interactions between HD2C and 14-3-3 kappa or nu were analysed in transientlytransformed Nicotiana benthamiana epidermal leaf cells using BiFC assays and confocalscanning microscopy four days after agroinfiltration (A, C). Kappa and nu homodimerformations were investigated in parallel (B, D). Confocal images of BiFC signals obtained bysingle optical section are shown in A to D (YFP, left panel).A, C: BiFC fluorescence indicated that 14-3-3 kappa-HD2C and 14-3-3 nu-HD2C interactions tookplace in nuclei (N) and nucleoli (No). The nucleolar signals were so strong that the excitationlight intensity had to be reduced to reveal the nucleolar structure.B, D: Kappa homodimers localised to the cytosol (Cyt) and the nucleus (N) but were absentfrom the nucleolus whilst nu homodimers localised mostly to the cytosol.The merged images of YFP and bright field images (YFP/BF) are shown for comparison (A to D,right panel). Scale bars = 50 μm.E: Interaction of HD2C with other 14-3-3 isoforms was confirmed (+, data not shown). Thelocalisation patterns were identical to those shown in A and C.

E

YFP Merged (YFP/BF)

kappa-YN/kappa-YCB

nu-YN/nu-YCD

nu-YN/HD2C-YCC

N

N

No

Cyt

N

kappa-YN/HD2C-YCA

N

No

Cyt

No



4.3. Discussion

4.3.1. 14-3-3 isoforms can dimerise freely with each other in vivo

In this project, dimeric interactions between ten out of thirteen expressed

Arabidopsis 14-3-3 isoforms and their intracellular localisations were visualised in

living plant cells using a transient BiFC assay. It was clearly shown that under the

experimental conditions used, all tested 14-3-3 isoforms can dimerise freely with each

other in vivo, suggesting that there is no obvious preferential dimerisation among

14-3-3 isoforms. These results verified previous reports, which were based on small

numbers of isoforms tested in vitro (Wu et al., 1997b; Abarca et al., 1999). The work

presented here further demonstrated that this freedom of dimerisation also applies to

the in planta situation. Published work using Arabidopsis (Wu et al., 1997b; Abarca

et al., 1999) and the results from this study are in contrast to those of mammalian and

yeast work which showed some degree of preferential dimerisation (Chaudhri et al.,

2003; Wilker et al., 2005). Thus, 14-3-3s in Arabidopsis may differ from their

mammalian and yeast counterparts; however, it remains to be investigated whether

this is the case for 14-3-3 proteins from other plants.

4.3.2. Subcellular localisation of 14-3-3 dimers

Published localisation studies using GFP fusion approaches demonstrated that

Arabidopsis 14-3-3 kappa, lambda, mu, omega, phi, and psi isoforms localised to the

cytosol and the nuclei in living plant cells (Cutler et al., 2000; Koroleva et al., 2005;

Paul et al., 2005; Rienties et al., 2005; Reumann et al., 2009; Table 4-1). The present

study using transient BiFC assays in Nicotiana leaf cells further verified that these

14-3-3 isoforms (not included the lambda isoform) were able to form homodimers

and any combination of heterodimers in those cellular compartments (Figs. 4-1, 4-2

and data not shown). Surprisingly, neither the published GFP-fusion approaches nor


the transient BiFC assays presented in this work showed chloroplastic localisation of

14-3-3 proteins. This is contrary to previous reports that 14-3-3 proteins were

detectable in the chloroplasts (Sehnke et al., 2000). Sehnke and co-workers showed

the presence of Arabidopsis 14-3-3 isoforms epsilon, mu, nu and upsilon in both

cytoplasmic and chloroplastic protein fractions but omega isoform was not found in

the stroma extract (Sehnke et al., 2000). The discrepancies of 14-3-3 organellar

localisations might be due to from the use of different plant and/or experimental

systems. It may further be that 14-3-3s indeed localise to the chloroplasts but that

the amount is too low to be detectable in the BiFC assays shown here. Furthermore,

lack of a chloroplast import signal in 14-3-3s still leaves the question of how 14-3-3s

gain access to the chloroplasts. Finally, it cannot be excluded that the published

work showing chloroplast localisation only shows association of 14-3-3s with

chloroplasts, i.e. that 14-3-3s were found not ‘in’ but attached to chloroplasts. This

would be supported by the findings shown here where 14-3-3s are often found

surrounding chloroplasts.

A more detailed discussion regarding the obtained localisation data of 14-3-3

dimers is in the following paragraphs.

4.3.3. Do nuclear localisation patterns reflect function of 14-3-3 dimers?

Subcellular localisation studies indicated BiFC fluorescence due to 14-3-3

dimerisation was found in the cytosol and nuclei of transiently transformed Nicotiana

leaf epidermal cells (Figs. 4-1 and 4-2). The fluorescence intensity in the cytosol did

not change significantly from dimer to dimer but varied for the nuclei both between

dimers and within one dimer combination. Based on the fluorescence intensity in

nuclei, subcellular distributions of 14-3-3 dimers were classified into three patterns,

those with higher nuclear than cytosolic signal intensities, those with nearly equal


intensities and those were the cytosolic signal appeared stronger than the nuclear

signal (Section 4.2.3.3).

The BiFC fluorescence intensity could be an indicator of the accumulation level

of the 14-3-3-BiFC dimeric complexes within cells. The mechanism of 14-3-3 protein

translocations into nuclei is still not clear. Classical nuclear localisation signals (NLS)

were not identified in any of the Arabidopsis 14-3-3 isoforms using the PredictNLS

online program (http://cubic.bioc.columbia.edu/cgi/var/nair/resonline.pl; data not

shown). One proposed mechanism is that subcellular localisation of 14-3-3s is

driven by interactions with target proteins (Paul et al., 2005). This is supported by

the finding that disturbances of 14-3-3 and target protein interactions lead to

localisation pattern changes of nuclear 14-3-3 kappa- and lambda-GFP fusion proteins

out of the nucleus in tricome cells of transgenic Arabidopsis plants (Paul et al., 2005).

One point which has to be taken into account is that the interaction with client

proteins in the 14-3-3 BiFC assays presented here may be disturbed or changed, either

due to the fusion protein nature or their overexpression. This may change

localisation patterns of 14-3-3 dimers. To verify the hypothesis that nuclear

localisation of 14-3-3s is driven by client interactions, one would have to show

interaction of the dimer and simultaneously interaction with the client. Such

approaches may be useful in elucidating dimerisation and client interaction in a

cellular and subcellular context.

The transient BiFC assays indicated that all tested 14-3-3 isoforms were able to

form dimers with each other. However, this only indicates the potential of all 14-3-3s

to freely dimerise. It has to be taken into account that a strong, non-native

promoter was used and that all 14-3-3s were expressed in a non Arabidopsis

environment and in leave tissues. These conditions may not reflect the true nature

of the 14-3-3s and may give a somewhat distorted view. Ideally, the results would be

http://cubic.bioc.columbia.edu/cgi/var/nair/resonline.pl


verified in Arabidopsis using native 14-3-3 promoters, which would allow for inclusion

of 14-3-3 expression profiles. A study aimed to verify such interactions in

Arabidopsis, although using the CaMV 35S promoter is presented in Chapter 5.

The transient BiFC assay in Nicotiana leaves indicates that Arabidiopsis 14-3-3

kappa isoform showed intense labelling in the nucleus as a homodimer and also when

present as a heterodimer in combination with most other 14-3-3 isoforms, except nu

and omega which commonly showed weaker nuclear BiFC signals in dimerisations

(Table 4-2 and Figs. 4-4 to 4-5). It also appears that kappa and nu can impose their

subcellular localisation preferences onto other 14-3-3s. For example, the psi isoform

shows Pattern 1 labelling when paired with kappa but demonstrates mainly Pattern 3

fluorescence when paired with nu (Fig. 4-5 and Table 4-2). The predominantly

nuclear labelling of 14-3-3 kappa is supported by a study that showed a similar

nuclear labelling pattern in tricome cells expressing 14-3-3 kappa-GFP fusion proteins

(Paul et al., 2005).

In contrast to the predominantly nuclear labelling pattern of 14-3-3

kappa-related dimers, the 14-3-3 nu isoform seemed to preferentially localise to the

cytosol, decreasing even the nuclear BiFC signal of kappa in a nu-kappa heterodimer

when compared to kappa homodimers. Furthermore, nu homodimers appeared to

be almost absent from the nucleus. Similarly, heterodimers of nu with omega and

iota appeared to be almost exclusively found in the cytosol (Fig. 4-5 and Table 4-2).

Different fluorescence intensity patterns between dimers and also for one dimer

combination of 14-3-3 dimers may reflect different cellular situations and

requirements. This is supported by findings showing that differential accumulation

of GFP-14-3-3 omega fusion proteins in nuclei of transgenic Arabidopsis hypocotyl

cells is responsive to the cell cycle state (Cutler et al., 2000). Commonly, the

GFP-omega fusion protein accumulated less in the nuclei, showing labelling similar to


the Pattern 3 described in the present study. However, the labelled fusion protein

accumulated to high levels in the nuclei just after completion of nuclear division,

appearing similar to Pattern 1 labelling in the present study. Shortly before

completion of cytokinesis, the fusion protein departed from nuclei (Cutler et al., 2000).

Although differential nuclear accumulation of 14-3-3 dimeric BiFC fluorescence was

detected in this study as well, it was unlikely due to the cell cycle regulation as

observed in Cutler et al. (2000) since the Nicotiana epidermal cells used here were

already highly differentiated. However, by analogy, the different patterns observed

for one dimer or pattern differences between dimers may reflect biological states of

so far unknown functions.

Interestingly, all the 14-3-3-BiFC dimeric complexes tested here displayed

fluorescence in the nucleoplasm but were absent from nucleoli. This contrasts with

an earlier study in which 14-3-3 kappa-GFP fusion proteins were reported to be

present in nucleoli (Paul et al., 2005). As there is no membrane barrier between the

nucleoplasm and nucleolus, the exclusion of 14-3-3 dimers from the latter is

intriguing.

4.3.4. Is the novel 14-3-3 localisation associated with the ER and with

wound induced cell death?

When investigating the subcellular localisation of 14-3-3-BiFC dimers in

epidermal cells, a distinct cytosolic pattern consisting of punctuate BiFC fluorescent

bodies (designated as Pattern 4) was observed very occasionally, suggesting that these

cytosolic structures do not reflect a common destination for 14-3-3 dimers. In

contrast this pattern may indicate special circumstances or cellular conditions. The

fading of the labelling also implies that 14-3-3 proteins might be responsive to an

event inducing this pattern. Furthermore, this pattern was observed for many


14-3-3 dimer combinations (Table 4-3) suggesting that the localisation response is not

isoform specific.

Observations of 14-3-3-BiFC fluorescence in epidermal peels indeed improved

the resolution and quality of the epifluorescence images and assisted greatly on

identification of subcellular distribution of the 14-3-3-BiFC dimeric complexes (Section

3.2.7). On the other hand, the procedure of epidermal peeling inflicts a degree of

mechanical stress onto the cells. Thus patterns observed for 14-3-3 dimers may

reflect cellular responses to wound effects. Several observations suggested that

wound-induced cell death was in progress in peeled epidermal cells. Firstly, there

was a dramatic decrease of the cell number showing fluorescence in the peels

compared to that in the original pre-peeled region in the intact leaves (Section 3.2.7).

Secondly, quick fading of BiFC fluorescence in epidermal peels when compared to

intact leaves may indicate cell death progression (data not shown). In contrast, BiFC

fluorescence could remain in leaf discs semi-immersed in water for 48 hours (data not

shown). Notably, the novel Pattern 4 fluorescence was mostly observed in peeled

epidermal samples but hardly detected in intact leaf samples, suggesting the pattern

would be a response to a factor associated with preparation of the peels (data not

shown).

The nature of this novel distribution pattern was further investigated in the

work presented in chapter 6.

4.3.5. Seven 14-3-3 isoforms can interact with histone deacetylase 2C

Interaction specificity of 14-3-3 isoforms with the target protein HD2C was

examined. The results indicated that all seven 14-3-3 isoforms tested were able to

interact with HD2C (Fig. 4-7), suggesting little specificity of 14-3-3 isoforms with

respect to the ability of binding to HD2C. In contrast, several in vitro binding assays


and application of the yeast two-hybrid system indicated binding specificities of

Arabidopsis 14-3-3 isoforms with some target proteins, such as nitrate reductase

(Bachmann et al., 1996b; Kanamaru et al., 1999), PM H+-ATPase (Rosenquist et al.,

2000; Alsterfjord et al., 2004) and phototropin 1 (Sullivan et al., 2009). This

introduces the question whether interaction specificities can be revealed using BiFC

assays. It may be that overexpression with or without mis-expression of the tested

proteins hinders such analysis. Furthermore, the stability of the formed YFP complex

may prevent specificity studies unless these specificities completely prevent binding

between the analysed proteins. Further investigations are needed to examine these

questions.

4.3.6. 14-3-3-HD2C interactions occurs in the nucleoplasm and nucleolus

All tested 14-3-3-HD2C interactions led to strong fluorescent signals in the

nucleus and even stronger labelling in the nucleolus. This was surprising as all dimer

combinations tested were absent from the nucleolus and some 14-3-3 homo- and

heterodimer combinations, namely those including 14-3-3 nu, did not preferentially

localise to the nucleus. It can be assumed that a relatively large amount of target

protein is available when expressing it under the control of the CaMV 35S promoter.

This amount may be sufficient to recruit 14-3-3 proteins to a specific location, namely

the nucleus and nucleolus, even if the 14-3-3 as a dimer shows less affinity for this

location. The absence of interaction between 14-3-3s and HD2C in the cytosol was

in contrast to experiments performed in mammalian systems where interactions of

14-3-3 proteins with HDACs were only found in the cytosol (Grozinger and Schreiber,

2000; Wang et al., 2000). This may point to a different regulation of the

plant-specific HD2 class of histone deacetylases when compared to the mammalian

and universal HDACs.


In previous studies, HD2C was identified in the nucleolar proteome (Pendle et al.,

2005). This is in agreement with the finding in this study. GFP fusion approaches

indicated that HD2C exclusively localises to the nucleolus but is not detectable in the

nucleoplasm and cytoplasm (Pendle et al., 2005; Sridha and Wu, 2006). This is only

in partial agreement with the current study in which HD2C interaction with 14-3-3s

was clearly found in the nucleus in addition to the nucleolus. Both studies agree on

absence of HD2C from the cytoplasm.

In 14-3-3-GFP fusion studies, dual localisation of 14-3-3s in the nucleus and

cytoplasm were reported for the isoforms kappa, lambda, mu, omega and phi but only

kappa was also detected in the nucleolus (Koroleva et al., 2005; Paul et al., 2005;

Rienties et al., 2005; Reumann et al., 2009 and summarised in Table 4-1). The

proteomic analysis of the Arabidopsis nucleolus which identifies HD2C as a nucleolar

protein does not identify any 14-3-3 proteins in the nucleolar proteome (Pendle et al.,

2005). Thus, the discovery that 14-3-3-HD2C interactions occur in the nucleolus as

demonstrated in this study, is a novel finding with respect to 14-3-3 localisation. This

brings up the question why 14-3-3 proteins are not found in the nucleolus when using

other approaches, including the 14-3-3 BiFC dimerisation assays performed in this

study (Section 4.2.3.1). One explanation may be the high expression levels achieved

using the CaMV 35S promoter to drive 14-3-3 and HD2C expression. These

expression levels possibly lead to high enough protein amounts to lead to

mis-localisation of binding partners. If this is the case, it may be that high HD2C

levels can recruit 14-3-3s to the nucleolus.

On the other hand, high amounts of 14-3-3s may recruit HD2C to the nucleus.

Why high 14-3-3 levels may not recruit HD2C to the cytoplasm may be explained with

a potential inability of the two proteins to interact in the cytosol. Interactions

between 14-3-3s and a target protein are often dependent on the phosphorylation of


the target. Thus, if interactions of 14-3-3s with HD2C are not possible in the

cytoplasm, it can be speculated that HD2C is non-phosphorylated before entering the

nucleus, i.e. after synthesis. This must be a short time period as HD2C-GFP fusions

were undetectable in the nucleoplasm (Pendle et al., 2005; Sridha and Wu, 2006).

When in the nucleus or nucleolus, HD2C, or at least some of the HD2C protein

molecules become phosphorylated and as a consequence, 14-3-3s can bind. It may

also be proposed that HD2C in the phosphorylated form is normally only present in

the nucleolus. This would be in agreement with the literature showing HD2C

localisation only in the nucleolar proteome and localisation of a HD2C-GFP fusion in

the nucleolus only (Pendle et al., 2005; Sridha and Wu, 2006). However, artificially

high amounts of 14-3-3s can be recruited to the phosphorylated, nucleolar HD2C.

Upon interaction, some of the 14-3-3-HD2C complex may move to the nucleus. This

would make the observed interaction of 14-3-3s with HD2C an experimental artefact.

Another explanation for the undetectable 14-3-3-HD2C interaction in the cytosol

would be that the HD2C does not localise to the cytosol or does so for a very short

period of time, i.e. it is translocated immediately to the nucleus when synthesised.

This is supported by the finding that HD2C-GFP fusion protein was also not detectable

in the cytosol (Pendle et al., 2005; Sridha and Wu, 2006). If as this case, there would

be no chance for cytosolic 14-3-3s to interact with HD2C in the cytoplasm.

More studies are clearly needed to answer the novel questions arising from this

work. These could include investigations into the impact of phosphorylation status

of HD2C on 14-3-3 binding. This could be achieved by generating a loss of

phosphorylation of HD2C mutant protein at potential 14-3-3 binding sites and then

testing for subcellular interaction of the HD2C mutant with 14-3-3 proteins using BiFC

assays.

Chapter 5. Analysis of 14-3-3 dimerisatons in transgenic Arabidopsis plants 125

Chapter 5

Analysis of 14-3-3 Dimerisation in

Transgenic Arabidopsis Plants


5.1. Introduction

5.1.1. Arabidopsis 14-3-3 isoforms freely dimerise with each other in a

heterologous plant system

As previously described (Section 4.1.1), only a limited number of plant 14-3-3

isoforms were shown to dimerise with each other by using mainly in vitro approaches

or the yeast two hybrid system (Wu et al., 1997b; Abarca et al., 1999). This study, by

focussing on in planta approaches has successfully demonstrated that ten of the

thirteen expressed Arabidopsis 14-3-3 isoforms can freely dimerise with each other

(Chapter 4). However, as these studies were performed in the heterologous plant

system Nicotiana benthamiana it remains unclear if similar results can be obtained in

the native plant system Arabidopsis thaliana. It cannot be excluded that the use of a

non-homologous system may impact on dimerisation results. Thus, it was decided

to verify a number of the interactions in Arabidopsis. Ideally Arabidopsis would have

been used to investigate all possible dimer combinations; however, previous studies

have shown that transient BiFC assays in Arabidopsis were not easy to perform and

lacked reliability (Chapter 3). Thus, the larger study was performed in Nicotiana as

described in Chapters 3 and 4, with follow-up studies using transgenic Arabidopsis as

described here.

Dimerisation of 14-3-3-YN and 14-3-3-YC fusion proteins in the native system

and in the presence of native 14-3-3s and their interacting proteins reflect, to a

degree, endogenous 14-3-3 behaviour. Dimerisation itself can be tested for in

analogy to the transient Nicotiana system using fluorescence microscopy.

Furthermore, tissue and developmental dependency of 14-3-3 dimer formation and

impact of endogenous or exogenous factors can be tested for using transgenic plants.

The outcomes of such analyses could provide more profound insights into the


function of 14-3-3 dimers than in vitro methods and studies performed in yeast and

non-homologous plant systems.

5.1.2. A literature review of 14-3-3 gene expression in Arabidopsis

All 14-3-3-split YFP constructs used in this study were expressed under the

control of the strong, widely active CaMV 35S promoter. Thus, foreseeable problems

with the evaluation of dimerisation results was the likelihood of different expression

patterns and higher expression levels of the introduced 14-3-3 BiFC transgenes

compared to endogenous 14-3-3s. Thus, endogenous 14-3-3 expression patterns

have to be taken into account when designing experiments and evaluating

dimerisation results.

To assist in this analysis, it was assumed that knowledge of endogenous 14-3-3

gene expression patterns in Arabidopsis would be helpful. Such knowledge could

help in identifying suitable 14-3-3-YN and 14-3-3-YC combinations for the genetic

generation of double transgenic plants and provide required background information

to evaluate dimerisation results.

Literature searches for the expression profiles of Arabidopsis 14-3-3 genes were

initially conducted. Alsterfjord and co-workers examined transcript abundance of

14-3-3 genes using Northern blot analysis and showed that 14-3-3 chi, lambda, mu

and epsilon are highly expressed in rosette leaves, followed by omega, phi, upsilon, nu

and kappa with lower but clearly detectable expression, and very low expression

levels of psi, omicron, iota and pi (Alsterfjord et al., 2004). In another report,

transcripts of chi, psi, lambda, kappa, mu, epsilon, omicron and pi were detected in

seven-day-old Arabidopsis seedlings (Cao et al. 2007). Furthermore, mu transcripts

were found in flowers, stems and leaves of mature plants (Kuromori and Yamamoto,

2000). Daugherty and co-workers monitored chi-promoter activity in transgenic


Arabidopsis plants using β-glucuronidase (GUS) as a reporter (Daugherty et al., 1996).

GUS activities were detected in flowers, roots, siliques, and imbibed seeds, suggesting

that the chi isoform was expressed in those tissues (Daugherty et al., 1996).

Rosenquist and co-workers reported that omicron was expressed in flowers, leaves

and roots of Arabidopsis plants, while iota was detectable exclusively in flowers and

expression of pi was not detectable in the tissues examined (Rosenquist et al., 2001).

It can be concluded from the literature that at least eight 14-3-3s are expressed in

seedlings (Cao et al. 2007), a minimum of nine in rosette leaves (Alsterfjord et al.,

2004) and that no less than four 14-3-3 genes are expressed in flowers (Daugherty et

al., 1996; Kuromori and Yamamoto, 2000; Rosenquist et al., 2001). Despite these

publications, the reported expression data of the Arabidopsis 14-3-3 family only cover

a limited number of tissues and in most cases are restricted to a small number of

these genes. To obtain a more complete picture, a comprehensive meta-profile

analysis based on publicly accessible microarray data was performed in this study

(Section 5.2.1).

Taken together, the aims of this chapter were:

(1) To analyse expression profiles of endogenous Arabidopsis 14-3-3 genes using

Genevestigator.

(2) To generate transgenic Arabidopsis plants harbouring 14-3-3-YN or -YC constructs

and to use those to generate double transgenic -YN/-YC plants by genetic crosses.

(3) To examine 14-3-3 dimerisation in double transgenic Arabidopsis plants in a whole

plant context.


5.2. Results

5.2.1. A large number of endogenous 14-3-3s are co-expressed in

Arabidopsis

Knowledge of endogenous 14-3-3 gene expression patterns in Arabidopsis

would assist this study in identifying suitable 14-3-3-YN and 14-3-3-YC combinations

for the genetic generation of double transgenic plants and providing background

information to evaluate 14-3-3 dimerisation results. To complement a literature

review of 14-3-3 gene expression in Arabidopsis (Section 5.1.2), a meta-profile

analysis of Arabidopsis 14-3-3 gene expression data was performed using

Genevestigator software (http://www.genevestigator.ethz.ch; Zimmermann et al.,

2004). Data sets from four thousand and seventy gene expression arrays

(Affymetrix® ATH1: 22k genechips) were used for this analysis. The “Anatomy tool”

and “Development tool” designed by Genevestigator were applied to analyse

expression profiles of eleven of the Arabidopsis 14-3-3 genes over tissues and

developmental stages. Genes encoding 14-3-3 isoforms chi (At4g09000) and phi

(At1g35160) were not included in this work of meta-profile analysis as the

oligonucleotides probe set used (probeset: 255079_s_at) was not specific enough to

differentiate between these two genes (Genevestigator). GRF14 (At1g22290) and

GRF15 (At2g10450) were not included in this analysis as they are considered to be

pseudogenes (Rosenquist et al., 2001; Chevalier et al., 2009).

A heat map of 14-3-3 expression profiles indicated that at least eight members,

omega, psi, upsilon, lambda, nu, kappa, mu and epsilon, were co-expressed in most of

the tissue categories (Fig. 5-1 A) and across different developmental stages (Fig. 5-1 B).

In contrast, omicron transcript was only found in tissues such as cotyledons, radicles,

adult rosette leaves, cauline leaves, lateral roots and the elongation zone of roots (Fig.

5-1 A). Furthermore, omicron expression was relatively low compared to that of

http://www.genevestigator.ethz.ch/


A B

Fig. 5-1. Tissue and development expression of Arabidopsis 14-3-3 genes –A Genevestigator heat map analysis.

The heat maps show tissue specific (A) and developmental (B) transcript abundance of elevenArabidopsis 14-3-3 genes. Data are based on the Affimetrix® ATH1:22k chip set and werecomputed using the meta-profile analysis tool of the online Genevestigator software(http://www.genevestigator.ethz.ch). Dark blue indicates higher and light blue lowerexpression levels. White indicates no detectable expression. For detailed descriptions of theanatomical ontology and of developmental stages, please refer to the Genevestigator website.The analysis shown above was verified in June 2009.


other 14-3-3 genes when analysed during developmental stages (Fig. 5-1 B). Still

further limited in tissue distribution were the expression of iota, which was detected

in pollen, and of pi only found expressed in the chalazal endosperm (Fig. 5-1 A), with

extremely low expression of these two genes in all developmental categories (Fig. 5-1

B).

From the meta-profile analysis and the literature study, it was concluded that a

large number of endogenous 14-3-3s are widely co-expressed in Arabidopsis with

relatively high expression levels.

5.2.2. Generation of transgenic Arabidopsis plants constitutively expressing

14-3-3-split YFP fusion proteins

The overall aim of the work presented in this chapter was the analysis of 14-3-3

dimerisation in the homologous plant system Arabidopsis. To do this, the first goal

was to generate a set of transgenic Arabidopsis plants each carrying either a

14-3-3-YN or a 14-3-3-YC construct by transforming wild-type (wt) plants (Col-0) using

the floral dip method (Section 2.4.5) with Agrobacteria carrying 14-3-3-split YFP

constructs. The constructs were chosen from the 14-3-3 library generated in the

work described in Chapter 3. The choice of clones was determined by their

availability during this project, the ability to match full-length with N-terminal deletion

mutant versions of 14-3-3s as these were to be used as negative controls, and the

expression patterns of 14-3-3 genes as determined from the literature and the above

described Genevestigator analysis.

For each transformation, 15,000 to 25,000 T1 seeds were screened for

resistance to either Basta or hygromycin, depending on the selection marker

contained on the transformed plasmid (Chapter 3). Wild-type plantlets were treated

in parallel with the selective agents to verify the expected sensitive phenotype.


Under both selective treatments, wt seedlings showed inhibited growth, lighter green

cotyledons and leaves, and died when kept under selective conditions. T1 seedlings

from transformed seed batches with wt-like phenotypes under selective conditions

were discarded as non-transgenic offspring (Fig. 5-2 and data not shown). Seedlings

with darker green cotyledons and leaves (under Basta selection; Fig. 5-2 B) or which

continued to grow (under hygromycin selection; Fig. 5-2 A) were selected as putative

transformants, transferred into individual pots with fresh compost mix and grown to

produce seeds. Presence of the transgene in the putative transformants was

confirmed by PCR analysis (Section 2.4.9) of genomic DNA from rosette leaves (data

not shown). Confirmed T1 plants were assigned individual line numbers (Table 5-1).

Fig. 5-2. Selection of transgenic Arabidopsis 14-3-3 BiFC seedlings.

Wild-type A. thaliana plants (Col-0) were transformed with 14-3-3-split YFP constructs using

the floral dip method. Two types of vectors were used, one conferring hygromycin and the

other Basta resistance (Chapter 3). T1 seeds were collected and germinated on either

hygromycin B (15 μg ml-1

) containing artificial media plates (A) or on soil for Basta (200 μM)

selection (B) as described (Section 2.4.7).

Seedlings were screened for resistance to the applied treatments 9 days (A) or 14 days (B)

after sowing. Hygromycin resistant seedlings grew beyond the germination stage; whilst the

growth of non-transformants was inhibited after germination (A). Seedlings resistant to Basta

stayed green and continued to grow after Basta application and were clearly distinguishable

from sensitive seedlings which appeared light to yellowish green and wilted (B). Arrows

indicate potential transformants.


Table 5-1. Summary of A. thaliana transformants carrying 14-3-3-split YFP

constructs.

Summarised are the result of Arabidopsis transformations with 14-3-3 BiFC constructs. Shown

are the names of the constructs used for transformation, the constructs, resistance by the

selectable plant transformation marker, names and numbers of isolated T1 plants, the ratios

of normal to aberrant phenotypes in the T1 generation and the names of plants with aberrant

phenotypes.

Construct

name 14-3-3-split YFP construct

Plant

selection*

Given name of

the T1 plants

Ratio of normal to

aberrant

phenotypes in the

T1 generation

Name of theT1

plants with aberrant

phenotypes

Ag3 pG nuX-229N-YN Basta Ag3-1 to 2 2/0 -

Ag4 pG nuX-229N-YC Basta Ag4-A1 toA2 2/0 -

Ag7 pG iotaXPCR-229N-YN Basta Ag7-1 1/0 -

Ag8 pG iotaXPCR-229N-YC Basta Ag8-1 to 4 4/0 -

Ag10 pG iota∆PCR-229N-YC Basta Ag10-1 to 6 6/0 -

Ag15 pG epsilonX-229N-YN Basta Ag15-1 1/0 -

Ag16 pG epsilonX-229N-YC Basta Ag16-1 1/0 -

Ag20 pG epsilon∆-179N-YC Hyg Ag20-1 0/1 20-1

Ag29 pG epsilon∆-229N-YN Basta Ag29-1 to 2 1/1 29-1

Ag22 pG omegaXPCR-229N-YC Basta Ag22-1 to 9 8/1 22-2

Ag23 pG omegaXPCR-179N-YN Hyg Ag23-1 to 8 8/0 -

Ag31 pG kappaXPCR-229N-YN Basta Ag31-1 to16 14/2 31-7, 31-16

Ag32 pG kappaXPCR-229N-YC Basta Ag32-1 to12 9/3 32-1, 32-9, 32-10

Ag33 pG kappa∆PCR-229N-YN Basta Ag33-1 to 2 2/0 -

Ag34 pG kappa∆PCR-229N-YC Basta Ag34-1 to 2 0/2 34-1, 34-2

Ag35 pG lambdaX-179N-YN Hyg B Ag35-1 to 4 4/0 -

Ag39 pG omicronXPCR-229N-YN Basta Ag39-1 to 11 9/2 39-3, 39-10

Ag40 pG omicronXPCR-229N-YC Basta Ag40-1 to11 9/2 40-2, 40-7

Ag50 pG muX-179NS-YC Hyg Ag50-1 1/0 -

Ag51 pG muX-179NS-YN Hyg Ag51-1 to 2 2/0 -

Ag60 pG mu∆-179NS-YN Hyg Ag60-1 to 2 1/1 60-2

Ag61 pG mu∆-179NS-YC Hyg Ag61-1 to 3 3/0 -

Ag53 pG phiX-179NS-YN Hyg Ag53-1 to 7 6/1 53-3

Ag54 pG phiX-179NS-YC Hyg Ag54-1 to 8 6/2 54-4, 54-8

Ag52 pG phi∆-179NS-YN Hyg Ag52-1 1/0 -

Ag55 pG psiX-179NS-YN Hyg Ag55-1 to 7 7/0 -

Ag56 pG psiX-179NS-YC Hyg Ag56-1 to 4 4/0 -

Ag59 pG psi∆-179NS-YC Hyg Ag59-1 1/0 -

Ag57 pG chiX-179NS-YN Hyg Ag57-1 1/0 -

Ag65 pG chiX-179NS-YC Hyg Ag65-1 1/0 -

subtotal 115 (normal) /18 (aberrant)

* Hyg, hygromycin B.


Thirty different BiFC constructs were successfully transformed into Arabidopsis,

including eleven full-length 14-3-3-YN constructs, ten full-length 14-3-3-YC constructs,

and additionally four 14-3-3∆-YN and five 14-3-3∆-YC constructs, which are

N-terminally truncated 14-3-3 mutant constructs (Table 5-1). A total of one hundred

and thirty-three independent T1 transgenic plants were obtained (Table 5-1). The

numbers of T1 plants isolated for the individual constructs ranged from one (e.g. for

pG iotaXPCR-229N-YN, Table 5-1) to sixteen (e.g. for pG kappaXPCR-229N-YN, Table

5-1) indicating that transgenic plants were obtained at a frequency of 0.005% - 0.1%;

well below published values of 0.5% - 3% for the floral-dip method (Clough and Bent,

1998). This was not further investigated as transgenic plants were obtained despite

these low frequencies.

5.2.3. Phenotypic changes were observed in some of the T1 transformants

Most of the T1 transgenic plants (115 out of 133) appeared similar to wild-type

plants (Table 5-1). The remaining eighteen T1 transformants showed aberrant

phenotypes, which may be related to the transgene itself, hinting at a biological

function, or caused by the insertion of the transgene into the genome which could

have an impact on function of the genes at the inserted loci. A differentiation

between possible causes was beyond the aims of this study. Hence, the observed

phenotypes, which are summarised in Table S1 (Appendix II), were solely used to

exclude transgenic plants with aberrant phenotypes from further analysis of 14-3-3

dimerisation.

5.2.4. Genetic analysis of T2 populations of 14-3-3-split YFP transformants

Timing and availability allowed for the genetic analyses of 16 out of the 133

obtained 14-3-3-split YFP transformants in the T2 generation (Table 5-2).


Table 5-2. Segregation of selectable markers in T2 progenies of sixteen

independent 14-3-3-split YFP transformants.

T2 populations of sixteen 14-3-3-split YFP carrying transgenic plants were tested for

resistance (R) and sensitivity (S) to selective agents (Basta or hygromycin; see also Table 5-1).

Segregation ratios were tested by χ2 test for fitness to possible segregation ratios. The null

hypothesis for χ2 tests was rejected at the 5 % level (P < 0.05). The expected ratios with the

highest probability of fitness were selected and are shown in the table. The χ2 test was not

applicable (n.a.) when any of the expected segregation frequency was less than 5 (Preacher,

2001).

Transformant

(genotype)

No. of T2

analysed

Resistant to

selection (R)

Sensitive to

selection (S) χ

2 test P-value

Ag3-1 (nu-YN) 16 9 7 1R:1S 0.62

Ag3-2 (nu-YN) 16 13 3 (3R:1S) n.a.

Ag4-A1 (nu-YC) 94 57 37 2R:1S 0.21

Ag4-A2 (nu-YC) 9 5 4 (1R:1S) n.a.

Ag15-1 (epsilon-YN) 22 17 5 3R:1S 0.81

Ag16-1 (epsilon-YC) 29 21 8 3R:1S 0.75

Ag29-2 (epsilon∆-YN) 23 13 10 1R:1S 0.53

Ag33-1 (kappa∆-YN) 22 14 8 2R:1S 0.76

Ag33-2 (kappa∆-YN) 20 16 4 3R:1S 0.61

Ag50-1 (mu-YC) 28 24 4 (11R:1S) n.a.

Ag51-1 (mu-YN) 23 23 0 All R -

Ag51-2 (mu-YN) 21 16 5 3R:1S 0.90

Ag52-1 (phi∆-YN) 21 21 0 All R -

Ag57-1 (chi-YN) 22 13 9 2R:1S 0.45

Ag59-1 (psi∆-YC) 10 10 0 All R -

Ag60-1 (mu∆-YN) 12 9 3 (3R:1S) n.a.

To gain an insight into the number of loci with a T-DNA insertion, segregation for

T-DNA linked hygromycin/Basta resistance was determined. The observed

segregation ratios were then statistically tested by chi-square (χ2) goodness-of-fit test

for fitness to different, theoretically expectable resistant to sensitive (R:S) segregation

ratios (Lu et al., 2004). The tested ratios included 1R:1S, 2R:1S, 3R:1S, 11R:1S and

15R:1S. The null hypothesis for the χ2 test was rejected at the less than 5% level


(P < 0.05). For each family analysed, the expected R:S ratio with the highest P-value

was selected and is listed in Table 5-2 to indicate the most likely segregation ratio

suggested by the observed frequencies. It should be noted that the use of the χ2

tests is inappropriate if any expected frequency is below 1 or if more than 20% of the

expected frequencies are less than 5 (Preacher, 2001).

Analysis of resistance to Basta for 16 Ag3-1 and 23 Ag29-2 T2 siblings revealed

segregation ratios of 9R:7S and 13R:10S. Both ratios fitted best a 1R:1S segregation

ratio with P=0.62 (Ag3-1) and P=0.53 (Ag29-2; Table 5-2) which is likely an indication

for insertion of a T-DNA at a single locus that led to gametophytic lethality in either

male or female gametes (Howden et al., 1998; Lu et al., 2004).

Four transformants, Ag15-1 (17R: 5S), Ag16-1 (21R:8S), Ag33-2 (16R: 4S) and

Ag51-2 (16R: 5S), appeared as a single locus T-DNA insertion lines that resulted in a

typical Mendelian segregation of 3R:1S ratio in the T2 populations.

Three transgenic lines, Ag4-A1, Ag33-1 and Ag57-1, exhibited a 2R:1S

segregation ratio indicating the potential of single locus inheritance with non-viable

homozygote plants (Lu et al., 2004).

The T2 seedlings of Ag51-1 (n=23), Ag52-1 (n=21) and Ag59-1 (n=10) examined

were all resistant to hygromycin, suggesting that there were at least two independent

T-DNA loci in the genome of these transformants.

The remaining transformants, Ag3-2, Ag4-A2, Ag50-1 and Ag60-1 had

segregation ratios for which the T2 populations were too small for an appropriate χ2

goodness-of-fit test (Table 5-2; n.a.) as more than 20% of the expected frequencies

were less than 5 (Preacher, 2001). Further genetic and molecular analyses are

needed to elucidate or confirm the number of T-DNA loci in the transgenic genomes.


5.2.5. Generation of “double” transgenic 14-3-3-YN and 14-3-3-YC

Arabidopsis plants for BiFC analysis of 14-3-3 dimerisation in a whole

plant context

The use of Nicotiana for transient BiFC analysis of Arabidopsis proteins has

proven to be a useful way of examining 14-3-3 dimerisation (Chapter 4). One

disadvantage with such an approach is the fact that the analysis was not done in the

native plant system. Additionally, transient BiFC analysis in Nicotiana was limited to

leaf tissue and was not extendable to a whole plant. In order to scrutinise the data

obtained in the transient analysis and to allow for whole plant dimerisation studies, it

was decided to perform BiFC investigations, limited to a few 14-3-3 dimer

combinations, in stable transformed Arabidopsis plants. To this end, a series of

transgenic Arabidopsis plants was generated harbouring one 14-3-3 BiFC construct

each (Section 5.2.2). Microarray data analysis (Section 5.2.1) and the published

literature (Section 5.1.2) indicated that a large number of 14-3-3 genes are widely

expressed, including chi, epsilon and mu. Taking microarray data and availability of

transformants into account, four double transgenic lines containing both a 14-3-3-YN

and a 14-3-3-YC construct were generated by crossing single transgenic T1 plants. T1

plants were usually used here to save time. Although T1 plants are heterozygous for

the transgenes, the dominance of the transgenes allowed for easy selection of double

transgenic offspring plants by testing for a combination of hygromycin and Basta

resistance (if applicable) and by monitoring for expected BiFC signals.

In detail, the maternal Ag50-1 plant (mu-YC, hygR; Table 5-1) was crossed with

the paternal Ag57-1 plant (chi-YN, hygR; Table 5-1) to generate offspring containing

both mu-YC and chi-YN transgenes (Table 5-3). Similarly, crosses of Ag15-1 x Ag50-1

were conducted to generate epsilon-YN/mu-YC plants (Table 5-3). Double transgenic

mu-YN/mu-YC plants were obtained by crossing of Ag50-1 x Ag51-1 and vice versa


Table 5-3. Generation of double transgenic 14-3-3-split YFP Arabidopsis

plants by genetic crosses.

Double transgenic Arabidopsis plants carrying the indicated 14-3-3-BiFC constructs were

generated by genetic crosses of single 14-3-3 YN and 14-3-3 YC parents. F1 seeds obtained

were germinated on MS media with the indicated selective conditions (refer to Table 5-1).

Seedlings resistant to the selection(s) were examined for BiFC fluorescence indicative of

14-3-3 dimerisation using fluorescent microscopy.

Cross (♀ x ♂)

F1 genotype

(note: both alleles are

heterozygous, unless

otherwise stated)

Plant selectionc

dNo. of F1

seeds

planted

F1 seedlings

resistant to

selection(s)

F1

seedlings

sensitive to

selection(s)

Resistant F1

seedlings

with BiFC

fluorescence

Ag15-1 x Ag50-1 epsilon-YN/mu-YC Basta and Hyg 33 4 29 4

Ag50-1 x Ag57-1 mu-YC/chi-YN Hyg 16 10 6 2

aAg50-1 x Ag51-1 mu-YC/mu-YN Hyg

a84 9 3 2

aAg51-1 x Ag50-1 mu-YN/mu-YC Hyg 40 34 6 2

Ag60-1-1b x

Ag61-1 mu∆-YN/mu∆-YC Hyg 20 20 0 0

a The genetic cross of Ag50-1 x Ag51-1 generated mostly shrunken seeds (72 out of 84), which did not germinate; while

the reciprocal cross (Ag51-1 x Ag50-1) produced normal seeds. b The maternal plant Ag60-1-1 used for the crosses with Ag61-1 pollen was a T2 line. A follow-up genotypic examination

on the F1 populations Ag60-1-1b x Ag61-1 suggested the maternal plant Ag60-1-1 was a transgenic homozygote

plant (see text). c Hyg, hygromycin B.

d No., number.

(Table 5-3). The crosses of Ag60-1-1 (a T2 plant) with Ag61-1 (a T1 plant) were done

to generate mu∆-YN/mu∆-YC plants, which served as a negative control for mu-mu

homodimerisation (Table 5-3).

5.2.5.1. Selection of BiFC expressing, double transgenic Arabidopsis plants

F1 seeds obtained from the genetic crosses were germinated on MS media with

appropriate selection to test seedlings for segregation of resistance versus sensitivity

to the plant selection (Table 5-3). Resistant seedlings were analysed for presence of

both transgenes by monitoring YFP fluorescence, which was indicative of 14-3-3

dimerisation (Table 5-3). Of the 33 epsilon-YN/mu-YC (Ag15-1 x Ag50-1) F1 seedlings

obtained, four were resistant to Basta and hygromycin and showed BiFC fluorescence

(Table 5-3). In analogy, two seedlings showed BiFC fluorescence in the offspring of


the mu-YC and chi-YN cross (Ag50-1 x Ag57-1, Table 5-3). The cross of Ag50-1 x

Ag51-1 produced mostly shrunken F1 seeds (72 seeds out of 84), which did not

germinate (data not shown). The remaining 12 seeds germinated, with two hygR

seedlings showing BiFC fluorescence, indicating formation of mu-YC/mu-YN dimers

(Table 5-3). The reciprocal cross of Ag51-1 x Ag50-1 was performed as well and it

produced normal seeds. Of the 40 Ag51-1 x Ag50-1 F1 seedlings, 34 were resistant

to hygromycin but only two showed BiFC fluorescence (Table 5-3).

Generation of mu∆-YN/mu∆-YC plants was achieved by crossing the maternal T2

plant Ag60-1-1 (mu∆-YN, hygR) with the T1 plant Ag61-1 (mu∆-YC, hygR). Twenty F1

seedlings tested were hygromycin resistance. Twelve of the F1 seedlings, randomly

selected to test for the presence of transgenes by PCR genotyping, contained the

mu∆-YN transgene with seven of those also containing the mu∆-YC transgene. This

suggested that the maternal plant Ag60-1-1 was homozygous and that the paternal

plant Ag61-1 was heterozygous, with a single locus for the transgene (mu∆-YC). As

expected, the seven F1 progeny containing both mu∆-YN and mu∆-YC transgenes did

not show BiFC fluorescence (Table 5-3 and data not shown).

5.2.5.2. Distribution of 14-3-3 dimers in double transgenic Arabidopsis

plants as observed by BiFC

An overview of the tissue distribution pattern of BiFC fluorescence in a young

chi-YN/mu-YC F1 seedling (Ag50-1 x Ag57-1, line 2) is shown in Figure 5-3 A to D.

Fluorescence intensity was strong in the root tip (Fig. 5-3 D), decreased in the

adjacent proximal zone (Fig. 5-3 D) and was strong again in the stele of the mature

root zone (Fig. 5-3 C). Fluorescence was also detectable in the hypocotyl and

cotyledons (Fig. 5-3 A, B). The distribution of BiFC fluorescence in epsilon-YN/mu-YC

and mu-YN/mu-YC seedlings resembled that of the seedling shown in Figure 5-3 (data

not shown). F1 seedlings exhibiting BiFC fluorescence were transferred to soil

Fig. 5-3. Localisation of a 14-3-3 mu-YC/14-3-3 chi-YN heterodimer as detected by BiFCanalysis of a genetically obtained double transgenic Arabidopsis F1 seedling.

Double transgenic 14-3-3 BiFC seedlings carrying two 14-3-3 BiFC constructs were obtained bygenetic crosses of single transgenic seedlings carrying 14-3-3 BiFC monomers. F1 seedlings wereexamined for BiFC fluorescence as an indicator of 14-3-3 dimerisation, using conventionalfluorescence microscopy. Shown is a seven day-old F1 seedling (Ag50-1 x Ag57-1 BiFC line 2,Table 5-3) carrying a 14-3-3 mu-YC and a 14-3-3 chi-YN constructs (mu-YC/chi-YN).Fluorescence indicating dimerisation of the two 14-3-3 constructs was observed throughout theseedling: cotyledons (A), hypocotyl and the junction of hypocotyl and root (B), root (C) and roottip (D). Strong fluorescence was observed in the root cap and the root stele with weak or nofluorescence in the adjacent proximal zone (D). Left panel: epfluorescence images (GFP filter).Middle panel: overlay of left and right panels. Right panel: bright field images. Scale bars =0.5 mm.


B

C

D

A mu-YC/chi-YN

Merged



and allowed to set seeds. The F2 progeny was used for a genetic analysis, a detailed

tissue specific dimerisation analysis and to monitor the subcellular distribution of

14-3-3 dimers in different tissues.

5.2.5.3. Genetic analysis of F2 offspring of double transgenic BiFC

Arabidopsis plants

F2 plants were derived by allowing the F1 plants shown in Table 5-4 to self

fertilise. The F2 seedlings were tested for hygromycin resistance and for BiFC

fluorescence. Out of 22 tested seedlings of Ag15-1 x Ag50-1 BiFC line1

(epsilonYN/mu-YC), 17 were resistant to hygromycin and 13 were BiFC positive (Table

5-4). The 17R:5S ratio (dominant hygromycin resistance) fitted a typical Mendelian

3R:1S segregation ratio with P=0.81 when tested using the chi-square goodness-of-fit

test (Section 5.2.4), suggesting that a single mu-YC allele (linked to hygromycin

resistance) was inherited from the Ag50-1 plant. BiFC fluorescence requires one

allele of each of the parents to be transmitted. The ratio of BiFC positive (13 BiFC+)

to BiFC negative (9 BiFC-) seedlings fitted an expected BiFC segregation ratio of 9

BiFC+:7 BiFC- with P=0.79, suggesting a single allele of the epsilon-YC transgene (linked

to Basta resistance) in the double transgenic parental plant.

In analogy, the observed hygromycin resistance ratio of 74R:6S in the offspring

of Ag51-1 x Ag50-1 BiFC line1 (mu-YN/mu-YC) fitted a 15R:1S ratio (P=0.64),

suggesting that the parental line contained two independent transgene alleles linked

to hygromycin resistance, one each from Ag51-1 and Ag50-1. This was further

supported by the observed BiFC segregation ratio (43 BiFC+:37 BiFC-; Table 5-4) which

fitted an expected 9 BiFC+:7 BiFC- ratio with P=0.65.


Table 5-4. Segregation of BiFC fluorescence in the F2 generation derived

from three independent double transgenic 14-3-3-split YFP Arabidopsis

lines.

The table shows segregation of hygromycin resistance and BiFC fluorescence (14-3-3

dimerisation) in the F2 progeny of double transgenic 14-3-3-split YFP plants. The F2 progeny

containing epsilon-YN/mu-YC were produced from one of the four BiFC-presenting lines

(Table 5-3) from the cross of Ag15-1 x Ag50-1. In analogy, the BiFC line 1 of Ag51-1 x Ag50-1

and the BiFC line 2 of Ag50-1 x Ag57-1 were used to generate F2 populations displaying BiFC

fluorescence of mu-YN/mu-YC and mu-YC/chi-YN dimerisation, respectively.

Transformant with BiFC Genotype Total *no. of F2

seeds planted

Resistant to

hygromycin

F2 Seedlings

with BiFC

BiFC line 1 of Ag15-1 x Ag50-1 epsilon-YN/mu-YC 22 17 13

BiFC line 1 of Ag51-1 x Ag50-1 mu-YN/mu-YC 80 74 43

BiFC line 2 of Ag50-1 x Ag57-1 mu-YC/chi-YN 116 107 53

* No., number.

In contrast, the segregation of hygromycin resistance to sensitivity of 11R:1S

(P=0.82) of the offspring of Ag50-1 x Ag57-1 BiFC line2 (mu-YC/chi-YN) indicated an

untypical Mendelian segregation of hygromycin resistance, suggesting two

independent T-DNA alleles at different, non-linked loci in the parental genome, with

one of the insertions leading to non-viable resistant homozygote offspring (Lu et al.,

2004). This was consistent with the conclusion drawn from a similar analysis

performed for the F2 generation of one of the parental lines, Ag57-1, with a 2R:1S

segregation ratio (Table 5-2). Segregation of BiFC fluorescence (53 BiFC+:63 BiFC-)

fitting a 1:1 ratio (P=0.35) rather than a 9 BiFC+:7 BiFC- ratio (P=0.02) further

supported this assumption (Table 5-4).


5.2.6. Distribution of 14-3-3 BiFC dimers in double transgenic Arabidopsis

plants

F2 plants in which fluorescence was detected were used to investigate the

subcellular localisation of the three 14-3-3 dimers in roots, cotyledon guard cells, and

epidermal cells of rosette leaves.

5.2.6.1. Distribution of 14-3-3 dimers in roots of transgenic seedlings

Five to nine day-old BiFC+ F2 seedlings were analysed for subcellular localisation

of 14-3-3 dimers in root tissues using confocal laser scanning microscopy (CLSM).

14-3-3 BiFC constructs were under the expression control of the CaMV 35S promoter.

Assuming that transcriptional regulation of the CaMV 35S promoter is independent of

the transgene itself, one can attribute differences in the distribution of the expressed

proteins to post-transcriptional and post-translational regulation, or to the impact of

the cellular environment on the ability of the proteins to interact. Comparing 14-3-3

dimerisation with expression of a control protein such as GFP, also under the control

of the CaMV 35S promoter, potentially shows promoter independent regulatory

events. Thus, obtained 14-3-3 dimerisation patterns were compared to fluorescence

observed due to CaMV 35S-GFP expression in control plants.

CLSM revealed strong GFP fluorescence in all zones, including meristematic and

elongation zones, of primary root tips of 35S-GFP seedlings (Fig. 5-4 B-D). GFP

fluorescence analysis thus demonstrated activity of the CaMV 35S promoter in all root

tip cells.

In contrast to GFP, strong BiFC fluorescence of mu-YC/chi-YN dimer complexes

was detected in the lateral and the columella root cap (Fig. 5-4 E-G). Very low levels

of fluorescence were observed in the meristematic and elongation zone of roots.

Fig. 5-4. Distribution of mu-YC/chi-YN and mu-YN/mu-YC dimeric complexes in root tips ofdouble transgenic Arabidopsis seedlings.

(A) Schematic illustration of root apical tissues delineating root cap (RC), meristematic (M) andelongation zone (EZ). Adapted from Marchant et. al., (1999) with permission from NaturePublishing Group, copyright© 1999.Confocal images of longitudinal section through the root tip of five to nine day-old transgenicArabidopsis seedlings expressing GFP (B-D), mu-YC/chi-YN fusion proteins (E-G; progeny ofAg50-1 x Ag57-1 BiFC line 2, Table 5-4) and mu-YN/mu-YC fusion proteins (H-J; progeny ofAg51-1 x Ag50-1 BiFC line 1, Table 5-4). GFP and 14-3-3-YN/YC expression was driven byCaMV35S promoters. BiFC fluorescence showed strong accumulation of 14-3-3 dimericcomplexes in the lateral and columella root cap. In contrast to GFP, 14-3-3 dimers were notobserved in meristematic and elongation zones of roots. Left panel (B, E and H): GFP or YFPfluorescence; right panel (D, G and J): the same root areas under bright field; middle panel (C, Fand I): overlay of left and right panels. Scale bars = 50 μm.

GFP

mu-YC/chi-YN

mu-YN/mu-YC

B

E

H

GFP/YFP Bright fieldMerged

A

C D

F G

I J



Similar 14-3-3 BiFC distribution patterns were also observed for mu-YN/mu-YC (Fig.

5-4 H-J) and epsilon-YN/mu-YC (data not shown) seedlings. Thus the differences in

GFP and BiFC fluorescence patterns suggested regulation events post transcription

initiation. For example, cellular conditions, e.g. native 14-3-3 binding competitors,

may interfere with efficient 14-3-3 dimerisation in cells or that post-transcriptional or

post-translational events prevent formation of the fusion proteins therefore showing

low level BiFC fluorescence.

Longitudinal optical sections imaged with CLSM were used to investigate the

distribution of epsilon-YN/mu-YC, mu-YC/chi-YN or mu-YN/mu-YC dimers or of GFP

fluorescence in epidermal/cortex cells of mature roots (Fig. 5-5). GFP fluorescence

was strong in the nuclei and cytoplasms of such cells (Fig. 5-5 A). Fluorescence of

epsilon-YN/mu-YC was clearly visible in the cytoplasm and nuclei, albeit fluorescence

in nuclei appeared weaker than in GFP plants (Fig. 5-5 D). Fluorescence in root cells

of mu-YC/chi-YN and mu-YN/mu-YC expressing plants was strong in the cytoplasm

with inconclusive results for nuclei of mu-YC/chi-YN plants and absent from nuclei of

mu-YN/mu-YC plants (Fig. 5-5 G and J).

5.2.6.2. Subcellular distribution of 14-3-3 dimers in cotyledon guard cells

In cotyledon guard cells, GFP fluorescence indicated that the GFP localised to

the nuclei and the cytoplasm but fluorescence was weak or absent from chloroplasts,

which appeared as red fluorescent structures when testing for chlorophyll

autofluorescence (Fig. 5-6 A and B). This was consistent with observations for GFP

localisation in root cells (Fig. 5-5 A). BiFC fluorescence of the 14-3-3

epsilon-YN/mu-YC dimeric complexes was detectable in the nuclei and the cytoplasm

with slightly lower intense in the nucleus (Fig 5-6 D), thus resembling Pattern 2 as

defined in transient expression experiments (Chapter 4). Fluorescence of the

Fig. 5-5. Subcellular localisation of 14-3-3-BiFC dimers in root cells of double transgenicArabidopsis seedlings.

Shown are confocal images of virtual longitudinal sections through a primary root of five totwelve day-old transgenic seedlings: (A-C) GFP, (D-F) epsilon-YN/mu-YC dimer (progeny ofAg15-1 x Ag50-1 BiFC line 1, Table 5-4), (G-I) mu-YC/chi-YN dimer (progeny of Ag50-1 x Ag57-1BiFC line 2, Table 5-4) and (J-L) mu-YN/mu-YC dimer (progeny of Ag51-1 x Ag50-1 BiFC line 1,Table 5-4). GFP expression was found in the cytoplasm and nuclei (A). All three 14-3-3 dimerswere found in the cytoplasm. Nuclear localisation was found for epsilon-YN/mu-YC, wasinconclusive for mu-YC/chi-YN and not found for mu-YN/mu-YC. Left panel: GFP (A) or YFPfluorescence (D, G and J); right panel (C, F, I and L), the same root areas under bright field;middle panel (B, E, H and K): overlay of left and right panels. Scale bars = 50 μm.

GFP/YFP Merged Bright field

A

GFP

G

mu-YC/chi-YN

D

epsilon-YN/mu-YC

J

mu-YN/mu-YC

N

B C

E F

H I

K L


Fig. 5-6. Subcellular localisation of 14-3-3-BiFC dimers in cotyledon guard cells of doubletransgenic Arabidopsis seedlings.

Shown are confocal images of single optical sections through cotyledon guard cells of nine totwelve day-old transgenic seedlings expressing GFP or two 14-3-3 BiFC constructs (A, D and G),merged images of GFP/YFP confocal images with red chlorophyll autofluorescence (Chl) (B, Eand H) and bright field images of the respective guard cells (C, F and I). Scale bars = 10 μm.(A-C) GFP fluorescence was detected in nuclei and in the cytoplasm of guard cells and wasweak or absent from chloroplasts.(D-F) YFP fluorescence, indicating the presence of epsilon-YN/mu-YC heterodimers in guardcells, was observed in the nuclei and cytoplasm but was undetectable or extremely low inchloroplasts.(G-I) YFP fluorescence, indicating the presence of mu-YN/mu-YC homodimers in guard cells,was strong in the cytoplasm, and very weak or absent from nuclei and chloroplasts.

D

GFP/YFP Bright fieldMerged with Chl

epsilon-YN/mu-YC

A

GFP

G

mu-YN/mu-YC

N

N

N

Chl

Chl

Chl

B C

E F

H I



mu-YN/mu-YC homodimer was clearly detectable in the cytoplasm but either absent

or only detectable at very low levels in nuclei (Fig. 5-6 G). Thus, the mu/mu

homodimer appeared in Pattern 3 as defined earlier (Chapter 4). The observations

made for the two 14-3-3 dimers in cotyledon guard cells matched those in root cells

of the same plants (Figs. 5-5 and 5-6).

It was reported that some 14-3-3 proteins, including epsilon and mu, localise to

chloroplasts (Sehnke et al., 2000). In the experiments performed here, fluorescence

was very week or absent from chloroplasts of guard cells for both the

epsilon-YN/mu-YC heterodimer and the mu-YN/mu-YC homodimer (Fig. 5-6 D-E and

G-H). Lack of BiFC fluorescence of 14-3-3 dimers in chloroplasts was consistent with

the observations made in transiently transformed Nicotiana leaves (Chapter 4).

Instead, fluorescence was clearly detectable surrounding the chloroplasts (Fig. 5-6 E

and H).

5.2.6.3. Subcellular distribution of 14-3-3 dimers in epidermal cells of

rosette leaves

The positive GFP control showed fluorescence in both the cytoplasm and the

nuclei of epidermal cells of rosette leaves of transgenic plants, with strong intensity in

the nuclei (Fig. 5-7 A-C).

Distribution of mu-YC/chi-YN, epsilon-YN/mu-YC and mu-YN/mu-YC dimerisation

was analysed in rosette leaves of transgenic Arabidopsis plants as well. As shown in

Fig. 5-7, both the mu-YC/chi-YN and epsilon-YN/mu-YC heterodimers were detectable

by BiFC fluorescence in the nuclei and cytosol (Fig. 5-7 D-F and G-I, respectively). In

contrast, mu homodimers (mu-YN/mu-YC) were detectable in the cytoplasm but not

in nuclei of the epidermal cells (Fig. 5-7 J-L). None of the three 14-3-3 dimers

Fig. 5-7. Subcellular localisation of 14-3-3-BiFC dimers in abaxial epidermal cells of rosetteleaves of double transgenic Arabidopsis seedlings.

Shown are confocal images of single optical sections through abaxial epidermal cells of rosetteleaves of nine to twelve day-old transgenic seedlings expressing GFP or two 14-3-3 BiFCconstructs (A, D, G and J), merged images of GFP/YFP confocal images with chlorophyllautofluorescence (B, E, H and K) and bright field images of the respective epidermal cells (C, F, Iand L). Scale bars = 50 μm.(A-C) GFP fluorescence was detected in nuclei (arrowheads) and in the cytoplasm of epidermalcells and was weak or absent from chloroplasts.(D-I) YFP fluorescence, indicating the presence of epsilon-YN/mu-YC (D-F) or mu-YC/chi-YN (G-I)heterodimers in epidermal cells, was observed in the nuclei (arrowheads) and cytoplasm butwas weak or absent from chloroplasts.(J-L) YFP fluorescence, indicating the presence of mu-YN/mu-YC homodimers in epidermal cells,was strong in the cytoplasm, but very weak or absent from nuclei and chloroplasts.

A

D

G

J

GFP/YFP Bright fieldMerged with Chl

GFP

mu-YC/chi-YN

epsilon-YN/mu-YC

mu-YN/mu-YC

B C

E F

H I

K L



appeared to localise to chloroplasts, which was consistent with the observations

made using guard cells (Fig. 5-6).

5.2.6.4. Distribution of 14-3-3 epsilon-YN/mu-YC dimer in flower tissues

The meta-profile analysis by Genevestigator indicating that 14-3-3 mu and

epsilon are expressed in flowers (Fig. 5-1) led to the decision to examine BiFC

fluorescence of 14-3-3 epsilon-YN/mu-YC dimers in flowers using conventional

fluorescence microscopy (Fig. 5-8). Analysis of negative control plants containing

only one 14-3-3 BiFC construct (14-3-3 phi-YC), which is unable to reconstitute a

functional YFP indicated strong autofluorescence in anthers (Fig. 5-8 B). Thus

anthers were excluded from observations described here.

Apart from anthers, fluorescence in the sepals, petals, stigma and in stamen

filaments confirmed that epsilon and mu, when co-expressed in flowers, are able to

dimerise with each other (Fig. 5-8 A). In sepals, petals and the stamen filaments,

fluorescence was prominent in the vascular tissues (Fig. 5-8 A, arrowheads).

5.3. Discussion

5.3.1. BiFC fluorescence in transgenic plants may reflect endogenous 14-3-3

dimerisations

Three sets of double transgenic 14-3-3 BiFC Arabidopsis plants,

epsilon-YN/mu-YC, mu-YC/chi-YN and mu-YN/mu-YC, were generated during this study.

It was foreseeable that the 14-3-3 BiFC constructs, driven by constitutive CaMV 35S

promoter, must be constitutively and ubiquitously expressed in the transgenic

Arabidopsis plants. On the other hand, Genevestigator microarray data (Fig. 5-1) and

reported transcript analyses (Section 5.1.2) indicated that a minimum of eight 14-3-3

isoforms (including epsilon and mu) are constitutively and ubiquitously expressed


Fig. 5-8. Distribution of epsilon-YN/mu-YC heterodimers in floral tissues of a doubletransgenic Arabidopsis plant.

This figure shows images obtained by conventional fluorescence microscopy through the GFPfilter set (left panel), the FITC filter set (middle panel) and bright field (right panel) of a doubletransgenic 14-3-3 epsilon-YN/mu-YC (Ag15-1 x Ag50-1, A) and a non-dimer forming, singletransgenic phi-YC control plant (Ag54-8, B). Fluorescence of 14-3-3 epsilon-YN/mu-YC wasdetected in most of the floral organs, including sepals (S), petals (P), stigma (St), carpel (C), andstamen filaments (F). Strong signals were observed in the vascular tissue of sepals, petals andstamen filaments (arrowheads). When these fluorescent signals were compared to the phi-YCcontrol plant (B), it became obvious that fluorescence in anthers (An) is, at least partially, dueto autofluorescence under the applied microscopic condition. Note, the white dotted linesshown in the left two panels outline partial margins of petals and sepals. Scale bars = 0.5 mm.

GFP filter Bright fieldFITC filter

A

B

epsilon-YN/mu-YC

phi-YC

St

An

P

S

C

F

C

S

PAn

F

St


and hence in most cell types will be co-expressed in a wt plant (Fig. 5-1). This leads

to the hypothesis that a number of native 14-3-3s, including the chi, epsilon and mu

analysed here, can be concurrently found across organs, tissues and cells. Thus,

combining 14-3-3 BiFC constructs under the regulatory control of the CaMV35S

promoter, although not identical to endogenous regulation, should not provide an

artificial situation with respect to protein expression and appeared suitable for an

initial dimerisation analysis. However, differences in expression levels between

native promoter and CaMV 35S promoter regulated expression of 14-3-3s as well as

addition of 14-3-3 expression on top of the native expression must be taken into

account when interpreting data. The BiFC fluorescence detected in the three sets of

double transgenic plants may be very close to the native situation for the three

respective 14-3-3 dimerisations: the heterodimers epsilon-mu and chi-mu and the mu

homodimer. Future experiments could be performed replacing the CaMV promoter

with endogenous 14-3-3 promoters and transformation of 14-3-3 knock-out lines to

ensure even closer resemblance to the native situation.

5.3.2. Visualising 14-3-3 dimerisations in a whole plant context

BiFC fluorescence was ubiquitously detectable in the three sets of double

transgenic 14-3-3 BiFC Arabidopsis plants (Figs. 5-3 to 5-8 and data not shown),

reflecting the fact that the three 14-3-3 dimerisations are able to occur across tissues.

Intriguingly, strong fluorescent signals of all the three 14-3-3 dimeric combinations

were observed in most of the root with the exception of the meristematic and

elongation zones of primary roots of seedlings (Fig. 5-4 and data not shown). In

these zones fluorescence was very weak or absent, possibly indicating a lack or low

concentrations of the 14-3-3-BiFC dimer molecules (Fig. 5-4).


BiFC fluorescence was compared to fluorescence due to a GFP construct

(pCAMBIA1302) expressed under the control of the CaMV 35S promoter. This

construct was clearly expressed in the meristematic and elongation zones of primary

roots as indicated by strong GFP fluorescence (Fig. 5-4 B-D). It was assumed that

expression patterns of the driven genes should be identical. However, when the

nucleotide sequences of 35S promoters used for the 14-3-3 BiFC constructs and the

GFP control were compared, it was found that they share ca. 600 bp identical

nucleotide sequences but that the promoter used in BiFC constructs was

approximately 200 bp longer (data not shown). It is unclear at this stage whether

region missing in the GFP linked 35S promoter contains repressive elements

preventing expression of 14-3-3 constructs in the meristematic and elongation zones

of roots. If this is not the case, then any difference in fluorescence distribution

between GFP and 14-3-3 BiFC in such zones may be not due to transcription level of

transgene expression but may indicate functional or biological impacts of cells on

14-3-3-BiFC dimer formation. The lack or low fluorescence may indicate that the

investigated 14-3-3s fail to dimerise in these areas or that post-transcriptional or

post-translational events prevent formation of the fusion proteins.

Expression of the 14-3-3- split YFP fusion proteins in such zones could be

verified by immunohistochemical analysis or by performing a western blot analysis on

proteins extracted from these zones using antibodies specific to the peptide tags

included in the fusion proteins. If protein expression could be confirmed, it would

be worthwhile investigating whether the 14-3-3 fusion proteins bind to target proteins

that modify the 14-3-3s or that prevent reconstitution of the split YFP fusion proteins

or even hinder 14-3-3 dimerisation. Protein pull-down assays have the potential to

demonstrate if the 14-3-3 fusion proteins bind to any target proteins in the tissue that

did not show BiFC fluorescence. A prevention of dimerisation in those root cells is


further supported as in all other tissues, fluorescence patterns of GFP and 14-3-3-BiFC

dimers appeared identical although their expressions were under the control of the

two 35S promoter variants.

Strong BiFC signals were detected in root caps and root steles for the three

dimer combinations tested (Figs. 5-3 and 5-4). This hinted that 14-3-3s play a role in

these tissues. Again, as the BiFC accumulation profiles of 14-3-3 dimers in these

tissues may only reflect the 35S promoter activity, further work should be performed

to verify this by replacing the 35S promoter with endogenous 14-3-3 promoters for

the BiFC analysis of 14-3-3 dimerisations.

5.3.3. Subcellular distribution of 14-3-3 dimer was somewhat

dimer-dependent in some tissues

On the subcellular level, localisation of 14-3-3 dimers appeared to be somewhat

dimer-dependent (Figs. 5-5 to 5-7). In epidermal cells of rosette leaves, the

epsilonYN/mu-YC dimer, as well as the mu-YC/chi-YN dimer, was found in both the

nucleus and cytoplasm (Fig. 5-7). In contrast, the mu-YN/mu-YC dimer localised

restrictedly to the cytoplasm, although nuclear staining such as DAPI stainingis

needed to confirm the nuclear position. Similar distributional differences for the

three 14-3-3 dimers were also observed in the seedling root cells and cotyledon guard

cells (Figs. 5-5 and 5-6, and data not shown). The distributional differences

suggested that the mu homodimer might play a role different from the heterodimers

epsilon-mu or chi-mu in these cells. One explanation for this is such dimers may

bind to different target proteins with different subcellular localisation as 14-3-3

localisation is postulated to be driven by client interactions (Paul et al., 2005).

Intriguingly, previous work has demonstrated Arabidopsis HD2C is able to interact

with several 14-3-3 isoforms, including chi, epsilon and mu, in the transiently


transformed Nicotiana leaves (Fig. 4-7). The HD2C-14-3-3 interactions localised

exclusively to the nucleus and prominently to the nucleolus (Fig. 4-7), demonstrating

the ability of HD2C to recruit 14-3-3 dimers, including the mu homodimer, into the

nucleus. Hence the absence of mu homodimeric BiFC from the nucleus of leaf

epidermal cells in transgenic Arabidopsis plants is contradictory to a possible

consequence of nuclear recruitment of 14-3-3 dimers by target proteins, at least by

endogenous HD2C, and therefore needs to be further investigated.

In contrast to the distributional differences detected in Arabidopsis leaf cells (Fig.

5-7), the three 14-3-3 dimers displayed similar localisation patterns in transiently

transformed Nicotiana leaf cells. In Nicotiana, the three 14-3-3 dimers all appeared

in nuclear localisation Patterns 1 or 2 (Figs. 4-1 and 4-5, Tables 4-2), i.e. stronger

fluorescence intensity in the nucleus than in the cytosol (Pattern 1) or similar intensity

in the nucleus and in the cytosol (Pattern 2). In leaf epidermal cells of stably

transformed Arabidopsis plants, however, nuclear BiFC signals were detected for both

the epsilon-mu and chi-mu dimers but not for the mu homodimer (Fig. 5-7).

Additionally, the fluorescent signals for both epsilon-mu and chi-mu dimers were

stronger in the cytosol than in the nuclei when observed by eye, thus neither

resembling Patterns 1 or 2 but instead Pattern 3 as previously defined in the Nicotiana

system (Chapter 4). From the results presented in this chapter, the outcomes of BiFC

performance of a given 14-3-3 dimer in the transient Nicotiana system were not

completely consistent with those from the Arabidopsis system. This supports a

common notion that functional analysis of a protein of interest in a heterologous

system should be eventually verified in the homologous system.

Chapter 6. Cell death induced aggregation and ER association of 14-3-3 dimers 156

Chapter 6

Cell Death Induced Aggregation

and ER Association of 14-3-3

Dimers


6.1. Introduction

When analysing the intracellular distribution of 14-3-3 dimers in epidermal cells

of transiently transformed Nicotiana leaves, a novel pattern was discovered during

this project (Pattern 4, chapter 4). This pattern, distinguished by small fluorescent

bodies, was detected for a number of 14-3-3 combinations but only observed

occasionally (Chapter 4, Table 4-3). The size and shape of these fluorescent bodies

were taken as an indication of organelle localisation of 14-3-3 dimers but may also

indicate aggregates of 14-3-3 proteins into protein bodies. As outlined in Section

1.6.2 (Chapter 1), 14-3-3 proteins were previously identified in chloroplasts (May and

Soll, 2000; Sehnke et al., 2000; Bunney et al., 2001) and mitochondria (Bunney et al.,

2001). Additionally, the Arabidopsis 14-3-3 isoforms lambda, mu, omega and psi

were identified in the proteome of peroxisomes (Table 4-1; Reumann et al., 2009).

Observations during this project could not confirm the presence of 14-3-3 dimers in

chloroplasts (Sections 3.2.7 and 4.2.3.2). This leaves mitochondria, peroxisomes, ER

or the Golgi as putative subcellular localisation of 14-3-3 dimers giving rise to the

novel distribution pattern.

The first aim of the work presented in this chapter was to determine if the novel

distribution pattern of 14-3-3 fluorescent bodies could be correlated with one of

these organelles (Section 6.2.1).

The fluorescent bodies observed upon 14-3-3 dimerisation appeared in cells

after performing epidermal peels and were rarely detected in intact leaf samples.

This suggested that a factor associated with the preparation of the peels, i.e.

wounding and possible cell death could induce this novel pattern. In animals,

14-3-3s are known as regulators of apoptosis (Fu et al., 2000; van Hemert et al., 2001;

Rosenquist, 2003). As in animals, regulated cell death is an essential process

occurring during plant development and when plants face stresses (reviewed in


Pennell and Lamb, 1997; Beers and McDowell, 2001; Jones, 2001; Lam, 2004). The

lack of plant homologues of animal apoptotic proteins suggests that plants may

employ a different regulatory mechanism for cell death than animals. It is well

established that transcription of plant 14-3-3 genes can be induced by some plant

pathogens and stress conditions (Roberts and Bowles, 1999; Lapointe et al., 2001;

Chen et al., 2006). However, an involvement of 14-3-3s in plant cell death is still not

clear (Rosenquist, 2003).

Two of the well-characterised examples of plant cell death are the development

of xylem elements and the hypersensitive response to pathogen attack (Lam, 2004).

Reactive oxygen species, such as hydrogen peroxide (H2O2), and stress hormones,

such as methyl jasmonate (MeJA), act as key signal molecules and/or effectors of the

cell death process (Beers and McDowell, 2001; Overmyer et al., 2003). Both

substrates were extensively used to trigger cell death in plant protoplast/cell culture

systems (Desikan et al., 1998; McCabe and Leaver, 2000; Houot et al., 2001; Paul and

Russell, 2001; Zhang and Xing, 2008).

The second aim of the work presented here was to investigate whether pattern

4 can also be induced in Arabidopsis thaliana and if cell death inducing conditions

other than wound induction can trigger this pattern. Double transgenic Arabidopsis

plants expressing two 14-3-3 BiFC constructs (Chapter 5) were used for these

investigations. Mesophyll protoplasts isolated from these Arabidopsis plants were

treated with exogenously applied H2O2 or MeJA and analysed for 14-3-3 dimer

patterns.

Thus, aims of the work presented in this chapter were:

(1) to determine if the novel distribution pattern of 14-3-3 fluorescent bodies were

correlated with intracellular organelles.


(2) To investigate whether the novel pattern of 14-3-3 dimers can also be induced by

cell death inducing conditions in Arabidopsis thaliana.

6.2. Results

6.2.1. Co-localisation of 14-3-3 dimers with fluorescent ER and Golgi

markers

Organelle-specific fluorescence dyes, such as MitoTracker® Red CMXRos

(Molecular Probes, Invitrogen) and ER-Tracker™ Blue-White DPX (Molecular Probes,

Invitrogen) were used to investigate if the fluorescent 14-3-3 BiFC bodies co-localise

with these organelles in epidermal peels of transiently transformed Nicotiana leaves.

Epidermal cells were examined for Pattern 4 by conventional fluorescence microscopy

(Section 2.5.1) and then stained with MitoTracker or ER-Tracker as per manufacturer’s

instruction. Dye concentrations up to the maximum recommended concentration (1

μM) for live cell staining were used, with staining times of up to one hour. However,

in all experiments, staining of individual organelles was unsuccessful and this

approach was consequently aborted (data not shown).

A second approach made use of co-expression of fluorescent proteins fused to

organelle targeting signals, which directed the fluorescent protein into specific

organelles. Co-localisation of BiFC fluorescence with that of the labelled protein

markers would indicate that 14-3-3 dimers were associated with a particular

organelle.

A peroxisomal targeted mRFP (Pracharoenwattana et al., 2005) and

mitochondrial, ER or Golgi targeted mCherry constructs (Nelson et al., 2007) were

chosen for the co-localisation analysis and allowed differentiation between the red

fluorescence of the organelle targeted markers and the yellow-green fluorescence of

14-3-3-BiFC complexes (Materials and Methods, Section 2.1.2.4). The organelle


marker genes were expressed under the control of a double 35S promoter

(Pracharoenwattana et al., 2005; Nelson et al., 2007) .

To allow for delivery into plant cells, plasmids containing the organelle marker

proteins were individually transformed into the same Agrobacterium strain (GV3101)

used for delivering 14-3-3-BiFC constructs into Nicotiana leaf cells (Section 2.3.4).

For co-localisation experiments, four Agrobacteria strains were co-injected into

Nicotiana leaves, one containing the organelle marker protein gene, one for each of

the two (YN and YC) 14-3-3 BiFC constructs and a fourth strain harbouring the p19

plasmid (Section 2.4.4). To identify the subcellular localisation observed as Pattern 4,

this work focussed on the 14-3-3 mu-epsilon heterodimer (mu-YN/epsilon-YC)

because Pattern 4 was more frequently observed with this dimer than with any of the

other 14-3-3 combinations (data not shown). Initially Nicotiana leaves were

infiltrated with a suspension mixture containing the four Agrobacterium strains in a

1:1:1:1 volume ratio. This resulted in very strong red fluorescence from the

expressed mCherry- or mRFP-tagged markers, impeding detection of BiFC

fluorescence caused by 14-3-3 dimerisation (data not shown). This may have been

due to higher marker protein expression as these were under the control of a double

35S promoter whilst 14-3-3 BiFC constructs were under the control of a single 35S

promoter. To overcome this problem, the amount of the marker-carrying

Agrobacterium strain in infiltration mixes was reduced 90% (i.e. to an 1:1:1:0.1

volume ratio). This led to the desired effect of lower marker expression and easier

detection of BiFC fluorescence, but also lowered the number of marker protein

expressing cells, a disadvantage which was accepted as co-localisation could be

investigated by observing a greater cell number.

To improve image resolution and to achieve easier comparison of organelle

marker and 14-3-3 dimer localisation, epifluorescence images were deconvolved using


Huygens Essential software (version 3.4, Scientific Volume Imaging B.V., The

Netherlands; http://www.svi.nl/). For comparison, the original and deconvolved

images are shown (Figs. 6-1, 6-3 and 6-4).

As described before (Chapter 4), the frequency of cells with Pattern 4 was very

low, between 1% and 3.3% cells. The low number of cells expressing the organelle

markers and the low frequency of Pattern 4 cells led, in most cases, to small numbers

of cells expressing the two together. Thus, the following results are based on small

numbers of cells, which needs to be taken into account when interpreting results.

a) Fluorescent bodies of mu-epsilon dimer were absent from

mitochondria

Only a small number of cells showed fluorescence of the mitochondrial marker

protein and 14-3-3 BiFC, and most demonstrated the 14-3-3 pattern previously

characterised as Pattern 2, i.e. 14-3-3 BiFC complex mediated fluorescence was found

evenly distributed between the cytosol and the nucleus (Chapter 4). In all cells

expressing the mitochondrial marker (transit peptide of ScCOX4 fused to mCherry;

Nelson et al., 2007), mitochondria appeared as small, red fluorescent bodies (Fig. 6-1

C and F), which localised to small, dark areas observed using the GFP filter set (Fig 6-1

A and D, position of dark areas indicated by arrowheads). BiFC fluorescence

indicating localisation of 14-3-3 mu-epsilon dimers was observed in the cytosol and in

a small number of cells with Pattern 4 in small vesicular structures (Fig. 6-1 A and D).

14-3-3 mediated BiFC fluorescence never co-localised with labelled mitochondria (Fig.

6-1 B and E). The different localisations of mCherry and BiFC labels were already

visible in non-deconvolved images (Fig. 6-1 B) but became much clearer after

deconvolution (Fig. 6-1 E). The results clearly demonstrated that 14-3-3 mu-epsilon

heterodimers were absent from mitochondria and that the punctate 14-3-3 BiFC

fluorescence described as Pattern 4 was in a different subcellular location.

http://www.svi.nl/


Fig. 6-1. Co-localisation analysis of 14-3-3 mu-epsilon heterodimeric BiFC complexes and amCherry-tagged mitochondrial marker in Nicotiana leaf epidermal cells.

N. benthamiana leaves were transiently co-transformed with an Agrobacterium mix containingbacteria harboring either 14-3-3 epsilon-YC, 14-3-3 mu-YN, p19 or an mCherry taggedmitochondrial marker gene. Figures A-C: four days after injection, leaves were analysed usingconventional fluorescence microscopy for expression and localisation of reconstituted YFP bydimerisation of mu-YN and epsilon-YC (A, green fluorescence) and the mitochondrial marker (C,red fluorescence). An overlay of A and C is shown in B. Figures D-F: the original epifluorescenceimages were deconvolved using Huygens Essential software to exclude out-of-focus information(Section 2.5.3). Using an RFP filter set, mitochondria were visible as small, red labeled, round tooval organelles which appeared as dark shadows under a GFP filter (arrowheads). In eachmicrograph, an area containing mitochondria and 14-3-3 dimers is shown enlarged (1.5x) forbetter resolution. The figures clearly showed that 14-3-3 BiFC dimers did not co-localise withmitochondria. Scale bars =10 μm.

GFP filter RFP filterMerged

Original

Deconvolved

A B C

D E F


b) Co-localisation analysis of fluorescent bodies of mu-epsilon dimer

and peroxisomes was inconclusive

Using the RFP-tagged peroxisomal marker (mRFP-PTS1; Pracharoenwattana et

al., 2005), peroxisomes were clearly visualised as round structures of similar size

(about 1.5 μm in diameter), widely distributed in transformed epidermal cells (Fig. 6-2

C). None of the cells investigated concurrently showed labelled peroxisomes and

BiFC fluorescent bodies in Pattern 4 of the mu-epsilon dimer. Instead, co-expressing

cells demonstrated Pattern 2 (Fig. 6-2 A). In such cells the peroxisomal marker and

14-3-3 dimers were not co-localised (Fig. 6-2 B). Thus, the results of investigating

14-3-3 dimers and labelled peroxisomes for co-localisation remained inconclusive.

Fig. 6-2. Co-localisation analysis of 14-3-3 mu-epsilon-BiFC complexes and a peroxisomal

mRFP marker in Nicotiana leaf epidermal cells.

Nicotiana leaves were transiently transformed with an Agrobacterium mix, as described in the

legend to Fig. 6-1, except a peroxisomal marker gene (mRFP-PTS1) replaced the mitochondrial

marker gene. Fluorescence in the transformed leaf epidermal cells was analysed four days

after injection as described (Fig. 6-1). Shown are images obtained for reconstituted YFP by

dimerisation of mu-YN and epsilon-YC (A, green fluorescence) and the peroxisomal marker (C,

red fluorescence). Overlay of GFP and RFP images (GFP/RFP) is shown in B and the merged

image of RFP and bright field (BF) is shown in C (RFP/BF). In the cell with a BiFC Pattern 2 (A),

14-3-3 mu-epsilon dimer did not co-localise with the peroxisome marker. None of the cells

investigated concurrently showed labelled peroxisomes and BiFC punctate bodies (Pattern 4)

of the mu-epsilon dimer. N, nucleus. Scale bars = 50 μm.


c) Fluorescent bodies of mu-epsilon dimer sometimes co-localise

with Golgi bodies

The expressed mCherry-Golgi marker (GmMan1-mCherry; Nelson et al., 2007)

localised to a large number of small, nearly round structures in transformed epidermal

cells (Fig. 6-3 C, F, I and L). As described for mitochondrial and peroxisomal marker

experiments, most of the cells showing concurrent BiFC fluorescence and red marker

protein fluorescence did not show the 14-3-3 Pattern 4, but instead 14-3-3 Pattern 2

(Fig. 6-3 A). In cells that did show 14-3-3 Pattern 4 and fluorescence of the Golgi

marker protein (Fig. 6-3 H and K), punctate BiFC fluorescence and red Golgi marker

fluorescence only partially co-localised (Fig. 6-3 K, overlap of fluorescence is indicated

by arrowheads).

d) Fluorescent bodies of mu-epsilon dimer co-localised partially

with ER-derivative structures

The ER can appear in at least two distinguishable conformations, reticular,

network like structures and vesicular structures. The reticular ER architecture was

easily detectable in epidermal cells of leaf discs from non-peeled Nicotiana leaves

transiently transformed with the ER-mCherry marker (signal peptide of

AtWAK2-mCherry-HDEL; Nelson et al., 2007; Fig. 6-4 C and F and Fig. 6-5). Within

such cells, BiFC fluorescence of 14-3-3 dimers localised within or associated with the

reticular structure as indicated by weak fluorescence (Fig. 6-4 A and B). Much

stronger BiFC fluorescence was observed in ’nodes‘ of the mCherry-labelled ER

network architecture (Fig. 6-4 B and E).

Fig. 6-3. Co-localisation analysis of 14-3-3 mu-epsilon heterodimeric BiFC complexes and amCherry-tagged Golgi marker in Nicotiana leaf epidermal cells.

Nicotiana leaves were transiently transformed with an Agrobacterium mix as described in thelegend to Fig. 6-1, except that a Golgi marker gene (GmMan1-mCherry) replaced themitochondrial marker gene. Fluorescence in the transformed leaf epidermal cells was analysed4 days after injection as described (Fig. 6-1). Shown are images obtained for reconstituted YFPby dimerisation of mu-YN and epsilon-YC (A and G, green fluorescence) and the Golgi marker (Cand I, red fluorescence). B shows the overlay of A and C and H shows the overlay of G and I.Figures D to F and J to L are deconvolved images of A to C and G to I, respectively. In cells withthe BiFC Pattern 2 (A to F), 14-3-3 mu-epsilon dimer did not co-localise with the Golgi marker.In contrast, the BiFC punctate bodies (Pattern 4) appeared to co-localise with some but not allstructures labelled by the Golgi marker (G to L, arrowheads). N, nucleus. Scale bars = 10 μm.

A

Original

Deconvolved

Original

Deconvolved

B C

D E F

G H I

J K L

N



Fig. 6-4. Co-localisation analysis of 14-3-3 mu-epsilon heterodimeric BiFC complexes and amCherry-tagged ER marker in Nicotiana leaf epidermal cells.

Nicotiana leaves were transiently transformed with an Agrobacterium mix, as described in thelegend to Fig. 6-1, except that an ER marker gene (signal peptide of AtWAK2-mCherry-HDEL)replaced the mitochondrial marker gene. Fluorescence in the transformed leaf epidermal cellswas analysed 4 days after injection as described (Fig. 6-1). Shown are images obtained forreconstituted YFP by dimerisation of mu-YN and epsilon-YC (A and G, green fluorescence) andthe ER marker (C and I, red fluorescence) in intact epidermal leaf discs (A to F) and in peeledepidermal cells (G to L). Overlays of A and C and G and I are shown in B and H, respectively.Figures D to F and J to L are deconvolved images of A to C and G to I, respectively. In epidermalcells of leaf discs, the ER-marker labeled a reticular architecture (C and F). 14-3-3 BiFCfluorescence was detected in discrete spots along this network (B and E, arrowheads). In peeledepidermal cells, BiFC fluorescence and mCherry fluorescence were closely associated.Unlabeled dark spots in G and J matched the ER marker labeled vesicular structures (compare Gand J with I and L), which were connected by a BiFC labeled network structure (H and K). Insetsin G-L (top, right): magnification (1.5x zoom) of the selected regions shown in G and J. Scalebars= 10 μm.

GFP filter RFP filterMerged

G

Original

Deconvolved

H I

K LJ

Original

Deconvolved

B CA

E FD


Better resolution of the ER network architecture was achieved by making a

composite image from a series of optical sections obtained by CLSM (Fig. 6-5 B). The

cell co-expressed the 14-3-3 mu-epsilon BiFC constructs in a network-like distribution

(Fig. 6-5 A). The identical localisation of 14-3-3 BiFC fluorescence and ER-mCherry

marker confirmed that 14-3-3 mu-epsilon dimers co-localised with the ER (Fig. 6-5 C).

In peeled epidermal cells, the ER-mCherry marker appeared in vesicle-like

structures (Fig. 6-4 I and L), and it was difficult to determine whether the ER marker

also labelled an ER network structure (Fig. 6-4 I). However, the absence of a

network-like structure in the deconvolved images suggested that the ER had

disintegrated into vesicular structures (Fig. 6-4 L). In Fig. 6-4 J-L BiFC fluorescence of

the 14-3-3 mu-epsilon dimer was clustered or associated with the red

fluorescence-labelled ER vesicular structures (Fig. 6-4 H and K).

In conclusion, the results of these analyses indicated that the 14-3-3 mu-epsilon

dimer did co-localise or associate with the ER. Some association was also observed

with the Golgi apparatus, but was completely absent from mitochondria. Small

vesicular structures appearing in leaf epidermal peels were shown to be of ER origin.

The ER derived vesicles were closely associated with 14-3-3 BiFC fluorescence as

observed in 14-3-3 Pattern 4.

6.2.2. Chemical cell death inducers and wounding trigger changes of

subcellular localisation of 14-3-3 dimers in transgenic Arabidopsis

The analysis of 14-3-3 dimerisation Pattern 4 in transiently transformed

Nicotiana cells was useful in revealing co-localisation of 14-3-3s with the ER.

However, as the 14-3-3 proteins are from Arabidopsis, Nicotiana is a heterologous

plant system. To investigate whether ER localisation can be observed in the

homologous Arabidopsis system, and to test conditions leading to cell death as


Fig. 6-5. Confocal image analysis of 14-3-3 mu-YN/epsilon-YC BiFC complexes and the ERmarker in transiently transformed Nicotiana leaf cells.

Confocal images were captured from cells co-transformed with 14-3-3 mu-YN/epsilon-YC BiFCconstructs and the ER targeting mCherry marker as described for Fig. 6-4.(A) A projection image of the YFP channel showed BiFC fluorescence indicative of 14-3-3 mu-epsilon dimerisation in the cytoplasm, the nucleus (N) and the ER.(B) A projection image of the mCherry channel showed the mCherry-labelled ER reticularstructure.(C) An overlay image of (A) and (B) demonstrated co-localisation of the BiFC and mCherryfluorescence, suggesting the 14-3-3 mu-epsilon dimer is associated with the ER.(D) A bright field image of the same field of view.Scale bars = 50 μm.

A B

C D

N


possible inducers of Pattern 4, double transgenic Arabidopsis plants expressing two

14-3-3 BiFC constructs (Chapter 5) were used. These were the same double

transgenic plants previously used to study 14-3-3 dimerisation in Arabidopsis, namely

epsilon-YN/mu-YC, chi-YN/mu-YC and mu-YN/mu-YC (Table 5-6).

6.2.2.1. The subcellular distribution of 14-3-3 dimers changes in response to

wounding in cotyledon epidermal cells

Cotyledons of 12-day-old mu-YN/mu-YC transgenic seedlings were wounded by

squeezing the tissue with tweezers. Homodimerisation of 14-3-3 mu was observed,

using CLSM by monitoring BiFC fluorescence in cells at the wounded sites one hour

after the treatment. As controls, cotyledons of non-wounded 14-3-3 BiFC plants

(negative controls) and wounded cotyledons of CaMV 35S-GFP plants (fluorescence

control) were analysed in the same way.

In non-wounded cells, fluorescence of mu-YN/mu-YC BiFC dimers was found

throughout the cytoplasm (Fig. 6-6 A). In wounded cells, fluorescence of 14-3-3 mu

BiFC complexes was visible in a larger area surrounding the vacuole, possibly

indicating that the vacuole had shrunken and that the cytosol took up more of the

cellular space (Fig. 6-6 B). Accumulation of 14-3-3 mu dimers was clearly visible in

the form of small round bodies distributed throughout the cytosol and sometimes

associated with a network-like structure, possibly cytoplasmic strands (Fig. 6-6 B,

arrowheads). The small bodies were similar in appearance to the bodies and the

ER-associated pattern observed in transient BiFC assays of peeled epidermal

Nicotiana cells (Figs. 4-6, 6-4 and 6-5).

As with 14-3-3 BiFC fluorescence, GFP fluorescence was observed in the cytosol

of non-wounded cells (Fig. 6-6 C). In addition, GFP fluorescence was observed in

nuclei. Upon wounding, vacuolar shrinkage was reflected by an extended area


Fig. 6-6. The subcellular distribution of 14-3-3-BiFC dimers changed upon wounding incotyledons of transgenic Arabidopsis seedlings.

Twelve-day-old cotyledons of transgenic Arabidopsis seedlings expressing 14-3-3 mu-YN/mu-YC(A and B) or GFP (C and D) were kept intact (A and C) or wounded by squeezing the tissues withtweezers tips (B and D) followed by confocal microscopy analysis one hour after treatments.Continuous, homogenous distribution of BiFC fluorescence was detected in the cytosol of non-wounded cells (A). In the wounded cells, the BiFC fluorescence of 14-3-3 mu homodimers wasvisible in a larger area surrounding the vacuole possibly indicating that the vacuole (V) hadshrunken and that the cytosol took up more of the cellular space (B). Accumulation of the 14-3-3 mu dimers was also clearly visible in form of small round bodies (arrowheads) distributedthroughout the cytosol and sometimes associated with a network-like structure, possiblycytoplasmic strands (B). GFP fluorescence was observed in both the cytosol and the nuclei ofnon-wounded cells (C). Upon wounding, vacuolar shrinkage was observable by an extendedarea showing GFP fluorescence (D). In contrast to 14-3-3 BiFC plants, wounding did not causethe formation of small fluorescent bodies in GFP plants. N, nucleus. Scale bars = 50 μm.

NN

C D

A B

Non-wounded Wounded

V


showing GFP fluorescence (Fig. 6-6 D). In contrast to 14-3-3 BiFC plants, wounding

did not cause the formation of small fluorescent bodies in GFP plants. The results

suggested that the observed wound-induced aggregations of BiFC fluorescence in

cells was 14-3-3 specific under the conditions of the experiment.

6.2.2.2. Subcellular distribution of 14-3-3 dimers in mesophyll protoplasts

derived from double transgenic Arabidopsis plants

To test whether the observed subcellular localisation Pattern 4 of 14-3-3 dimers

is associated with cell death and can be induced, a protoplast system was employed.

This has the advantage that all cells will be equally exposed to the treatment (McCabe

and Leaver, 2000), which is not guaranteed in a tissue context.

Protoplasts were isolated from rosette leaves of three transgenic 14-3-3 BiFC

lines, epsilon-YN/mu-YC, chi-YN/mu-YC and mu-YN/mu-YC (Section 2.4.3). Before

commencing with treatments, a CLSM analysis was performed to identify the

localisation of 14-3-3 BiFC dimers in protoplasts of the three transgenic lines.

BiFC fluorescence, indicative of 14-3-3 dimerisation, was found in the cytosol and

nucleus of protoplasts expressing epsilon-YN/mu-YC or mu-YC/chi-YN dimers but was

absent from chloroplasts in both (Fig. 6-7 A-D). The dual localisation of these two

14-3-3 heterodimers in both the nucleus and cytosol was in agreement with earlier

observations of the two dimers in rosette leaf epidermal cells of transgenic

Arabidopsis plants (Fig 5-7 D-F and G-I). However, a higher level of nuclear

accumulations of the two dimers were observed in mesophyll protoplasts (Fig. 6-7 A-D)

than those in leaf epidermal cells (Fig. 5-7 D-I).

Fluorescence of the mu-YN/mu-YC homodimer was also observed in the cytosol

and in the nuclei of mesophyll protoplasts (Fig. 6-7 E-F). This observation differed

from those made for this dimer in leaves in that nuclear localisation was not detected


Fig. 6-7. Subcellular localisation of 14-3-3 dimers as detected by BiFC fluorescence in rosetteleaf protoplasts derived from transgenic Arabidopsis plants.

Mesophyll protoplasts were obtained from rosette leaves of transgenic Arabidopsis plantsexpressing the following 14-3-3 BiFC dimer constructs: epsilon-YN/mu-YC (A-B), mu-YC/chi-YN(C-D) and mu-YN/mu-YC (E-F). A, C and E show single optical sections and B, D and F showprojection images of several serial optical sections. The panels from left to right show: YFPfluorescence (YFP), chlorophyll autofluorescence (Chl), merged images of YFP and Chl (YFP/Chl)and the overlay of YFP/Chl and a bright field image (YFP/Chl/BF).Dual localisation of BiFC fluorescence in both cytosol and the nucleus (N) was observed for allthe three 14-3-3 dimeric combinations. In contrast, BiFC fluorescence was not observed inchloroplasts for either epsilon-YN/mu-YC (A-B) or chi-YN/mu-YC (C-D), but was detected inchloroplasts for mu-YN/mu-YC dimer (E-F). Scale bars = 10 μm.

N

E

F

YFP YFP/ChlChl YFP/Chl/BF

N

N

C

D

mu-YN/mu-YC

N

N

A

B

N

mu-YC/chi-YN

epsilon-YN/mu-YC


in Arabidopsis leaf epidermal cells (Fig. 5-7 J-L). On the other hand, this dimer was

shown to localise to nuclei when analysed in transiently transformed Nicotiana leaves

(Chapter 4). In addition to nuclear and cytosolic localisation, this dimer was also

detectable in chloroplasts (Fig. 6-7 E-F). Again, this was in contrast to previous

observations made using leaves of transgenic Arabidopsis plants (Fig. 5-7 J-L) and

transiently transformed Nicotiana leaves (Sectioin 4.2.3.2), where this dimer appeared

absent from chloroplasts. Overall, with the exception of chloroplastic localisation of

the mu homodimer, localisation of 14-3-3 dimers in protoplasts agreed well with

localisation observed in leaves of transgenic Arabidopsis plants and transiently

transformed Nicotiana leaves.

6.2.2.3. Hydrogen peroxide treatment induces the novel subcellular

localisation of 14-3-3 dimers

Hydrogen peroxide was shown in a number of experiments to induce plant cell

death (Desikan et al., 1998; McCabe and Leaver, 2000; Houot et al., 2001; Paul and

Russell, 2001). Hence, H2O2 was used in the pilot study of testing the impact of

induced cell death on the distribution of 14-3-3 dimers (Section 2.4.10.1).

Treatment of GFP-expressing and 14-3-3 dimeric BiFC presenting protoplasts

with H2O2 for 24 h caused a dramatic loss of fluorescence in almost all protoplasts and

disintegration of cellular integrity regardless of the H2O2 concentration used (data not

shown). In contrast, most GFP-expressing and 14-3-3-BiFC protoplasts from the

non-H2O2-treated control (mock treatment) remained fluorescent and structurally

intact after 24 h incubation (data not shown). When incubated in the presence of

0.2 mM H2O2 for 12 h, a significant number of GFP-expressing and 14-3-3 protoplasts

had lost fluorescence and cellular integrity (data not shown); however, a sufficient

number remained intact for analysis. Protoplasts expressing 14-3-3

epsilon-YN/mu-YC BiFC dimers showed the formation of small, round, fluorescent


structures (Fig. 6-8) resembling those previously observed for 14-3-3 BiFC Pattern 4 in

transiently transformed Nicotiana epidermal peels (Figs. 4-6, 6-1, 6-3 and 6-4). In

contrast, no significant change of fluorescence patterns was observed in the

protoplasts expressing either chi-YN/mu-YC mu-YN/mu-YC or GFP when treated with

H2O2 (Fig. 6-8 and data not shown).

6.2.2.4. Methyl jasmonate treatment induces the novel subcellular

localisation of 14-3-3 dimers

The plant hormone MeJA is another widely used inducer of plant cell death

(Asai et al., 2000; Repka et al., 2004; Zhang and Xing, 2008). It was decided to test if

the novel distribution pattern of aggregates of 14-3-3 dimers was inducible by

exogenous MeJA.

Most 14-3-3 dimeric BiFC presenting protoplasts did not display significant

changes in the subcellular distribution of fluorescence when treated with MeJA (Fig.

6-9 and data not shown). The exception was a small number (ca. 1%) of

epsilon-YN/mu-YC protoplasts treated with 50 μM MeJA which displayed fluorescent

bodies (Fig. 6-9) similar to those observed as Pattern 4 (Fig. 4-6) and in hydrogen

peroxide treatments (Fig. 6-8 and data not shown). Such bodies did not appear in

mock-treated controls for the epsilon-YN/mu-YC protoplasts nor in the GFP controls

or in the chi-YN/mu-YC protoplasts (Fig. 6-9).

Effects of exogenous MeJA on subcellular localisation of the 14-3-3-BiFC dimers

were further examined in excised leaves. No significant changes on 14-3-3 BiFC or

GFP fluorescence distribution were detected in excised rosette leaves from

six-to-seven-week-old transgenic plants expressing either epsilon-YN/mu-YC,

c h i - Y N / m u - Y C , m u - Y N / m u - Y C , m u Δ - Y N/ m u Δ - Y C o r G F P c o n s t r u c t s


Fig. 6-8. Changes of epsilon-YN/mu-YC BiFC complex localisation were induced by H2O2

treatment in mesophyll protoplasts of transgenic Arabidopsis plants.

Mesophyll protoplasts were obtained from rosette leaves of transgenic Arabidopsis plantsexpressing a GFP construct (top row) or the epsilon-YN/mu-YC BiFC dimer (bottom row).Protoplasts shown in the left two columns were subjected to mock treatment (no H2O2).Protoplasts shown in the two right columns were treated with 0.2 mM H2O2 for 12 hours priorto analysis by conventional fluorescence microscopy. Distribution of GFP fluorescence wasunaffected by H2O2 treatment when compared to mock treated protoplasts. In contrast, BiFCfluorescence of epsilon-YN/mu-YC dimers aggregated into punctate bodies (arrowhead) afterthe H2O2 treatment. Scale bars = 50 μm.

Control (0 mM H2O2)

GFP

epsilon-YN/mu-YC

0.2 mM H2O2

GFP filter GFP filterBright field Bright field

GFP

epsilon-YN/mu-YC


Control 50 μM MeJA

Fig. 6-9. Changes of epsilon-YN/mu-YC BiFC complex localisation were induced in a smallnumber of mesophyll protoplasts of transgenic Arabidopsis plants by MeJA treatment.

Mesophyll protoplasts were obtained from rosette leaves of transgenic Arabidopsis plantsexpressing a GFP construct (top row), the chi-YN/mu-YC BiFC dimer (second row) or the epsilon-YN/mu-YC BiFC dimer (bottom two rows). Protoplasts shown in the left two columns weresubjected to mock treatment (0.1% ethanol). Protoplasts shown in the two right columns weretreated with 50 μM MeJA in 0.1% ethanol for 24 hours prior to analysis by conventionalfluorescent microscopy. Columns two and four show bright field images or bright field imagescombined with fluorescent images (column 4, row three). No apparent changes in subcellulardistribution were detected for GFP or the chi-YN/mu-YC BiFC dimer after MeJA treatment whencompared to mock treatments. In contrast, MeJA treatment caused aggregation of epsilon-YN/mu-YC BiFC complexes into punctate bodies (arrowheads) in a small number of protoplasts,two of which are shown here. This aggregation pattern was not observed in mock treatedprotoplasts (controls). The punctate bodies resembled the pattern 4 observed for 14-3-3 dimersafter wounding in Nicotiana and Arabidopsis leaves (Chapter 4 and Fig. 6-6 B, respectively).Note, the MeJA treated protoplast shown in row three possibly displays shrinkage of the cytosol.Scale bars = 50 μm, unless otherwise indicated.

GFP

mu-YC/chi-YN

epsilon-YN/mu-YC

25 μm

25 μm

25 μm

25 μm

GFP filter

GFP filter

Bright field

Bright field

epsilon-YN/mu-YC

epsilon-YN/mu-YC

mu-YC/chi-YN

GFP


treated with 50 μM MeJA for 38 h (Fig. 6-10). Fluorescence was absent from the

muΔ-YN/muΔ-YC negative dimerisation control (Fig. 6-10).

Methyl jasmonate inhibits root growth (Staswick et al., 1992). As 14-3-3 BiFC

plants are 14-3-3 overexpressors, testing MeJA on growth of such seedlings would

show: (1) if 14-3-3 plays a role in MeJA induced root growth inhibition and (2) if MeJA

has any impact on 14-3-3 dimer formation and localisation in newly grown root and

leaf tissues.

Ten day-old transgenic seedlings expressing GFP or BiFC fluorescence were

transferred from hygromycin selection MS plates to 0 μM or 50 μM MeJA containing

MS plates and allowed to grow for four more days. Absolute root growth during this

period was measured. As shown in Fig. 6-11, root growth inhibition by MeJA was

very similar in double transgenic 14-3-3 seedlings to that in the GFP control. This

suggested that overexpression of the tested 14-3-3-BiFC fusion proteins has no

impact on MeJA signalling with respect to root growth inhibition. Additionally, BiFC

fluorescence pattern was examined in the first pair of leaves and in root tips of

primary roots at the end of treatments. No significant changes were observed in

terms of 14-3-3 dimer formation and subcellular localisation between MeJA treated

and untreated seedlings within the newly grown tissues (Fig. 6-11).

Taken together, an altered distribution pattern of 14-3-3 dimers was detected in

cells after wounding and H2O2 treatment and to a lesser extent in MeJA treated

protoplasts. The observed pattern changes resembled those described for

epidermal peels from Nicotiana leaves. These results further suggested that the

novel 14-3-3 dimerisation Pattern 4 occurred in response to cell death inducing

conditions.


Control 50 μM MeJA

GFP filter Bright field GFP filter Bright field

GFP

chi-YN/mu-YC

epsilon-YN/mu-YC

mu-YN/mu-YC

muΔ-YN/muΔ-YC

Fig. 6-10. MeJA did not impact on subcellular 14-3-3 BiFC dimer distribution in intactepidermal cells of transgenic Arabidopsis rosette leaves.

Rosette leaves were excised from six-to-seven-week-old transgenic Arabidopsis plantsexpressing the indicated constructs and were subjected to mock treatment (0.1% ethanol) or50 μM MeJA in 0.1% ethanol. Effects of MeJA on the distribution of GFP or 14-3-3 BiFCfluorescence were compared to mock treatments 38 hours post treatment by conventionalfluorescent microscopy. No changes in distribution were detected. Scale bar= 50 μm.


Fig. 6-11. MeJA inhibited root growth, but did not affect either dimer formation ordistribution pattern of 14-3-3 dimers in newly grown leaf and root tissues.

Ten-day-old transgenic Arabidopsis seedlings expressing the indicated constructs weretransferred from hygromycin selection plates to 0 μM or 50 μM MeJA containing plates andallowed to grow for four days. Absolute root growth during this period was measured and theepifluorescence images were obtained from newly grown tissues (first pair of leaves or primaryroot tips) at the end of treatment. Root growth values are the means ± SD from 7 to 9seedlings as indicated. No significant changes were observed in terms of 14-3-3 dimerformation and tissue/cellular distribution between MeJA treated and untreated seedlingswithin the newly grown tissues. Scale bars for leaf and root samples are 10 μm and 100 μm,respectively.

0

10

20

30

Ctrl MeJA

mm mu-YN/mu-YC (n=7)

roo

t gr

ow

th

0

10

20

30

Ctrl MeJA

mm mu-YC/chi-YN (n=8)

roo

t gr

ow

th

0

10

20

30

Ctrl MeJA

mm epsilon-YN/mu-YC (n=9)

roo

t gr

ow

th0

10

20

30

Ctrl MeJA

mm GFP (n=8)

Ctrl MeJA

roo

t gr

ow

th

10 μm

100 μm

leaf

root


6.3. Discussion

6.3.1. The potential association with the ER upon cell death induction and

wounding suggest a role of 14-3-3s in ER mediated apoptosis

During identification of subcellular localisation of 14-3-3 dimerisations using

BiFC assay, a novel distribution pattern of 14-3-3 dimers, characterised as small,

punctate, cytoplasmic bodies (designated as Pattern 4), was observed in the cells of

epidermal peels of transiently transformed Nicotiana leaves (Chapter 4).

Co-expression of an ER targeting fluorescent protein marker with 14-3-3

mu-YN/epsilon-YC BiFC constructs indicated that the 14-3-3 mu-epsilon dimer

co-localised to ER reticular network (Fig. 6-5) and ER-derived structures (Fig. 6-4 A-F)

in epidermal cells of transiently transformed Nicotiana leaves. Sometimes, after

wounding caused by epidermal peeling, aggregation of this dimer was observed in

association with the Golgi apparatus (Fig. 6-3 G-L) and with very bright fluorescence in

punctate areas of the ER (Fig. 6-4 G-L). These preliminary observations have been

supported by more recent work in the Martin lab, which has shown that the network

structure with 14-3-3 BiFC fluorescence is an early event and that punctate bodies

appear at later stages after wounding or herbicide treatments (R. Li, unpublished

results).

The punctate areas in which 14-3-3 dimers localised to or associated with the ER

may be ER export sites/exit sites (ERES). The ERES are the start of transport vesicles

leaving the ER known as coatomer protein complex II (COPII) vesicles (Hanton et al.,

2005; Watson and Stephens, 2005; Budnik and Stephens, 2009). 14-3-3 regulation

on export of membrane proteins from ER has been studied in depth in yeast and

mammalian systems (Nufer and Hauri, 2003; Mrowiec and Schwappach, 2006;

Shikano et al., 2006). A proposed model for mammalian 14-3-3 regulated trafficking

of membrane protein targets suggests that binding to 14-3-3 reduces ER retention,


therefore promoting forward transport of membrane proteins to the plasma

membrane (Nufer and Hauri, 2003; Shikano et al., 2006). However, it is not known if

14-3-3s regulate protein export from the ER in plants.

Aggregation of 14-3-3 dimer into punctuate bodies was also observed for the

mu homodimer at wounded sites of cotyledons (Fig. 6-6) and for the epsilon-mu

heterodimer in H2O2- and less frequently in MeJA-treated mesophyll protoplasts (Figs.

6-8 and 6-9) of transgenic Arabidopsis 14-3-3 BiFC plants. The results strongly

support an idea that the morphological change of 14-3-3 dimer distribution was

responsive to cell death inducing treatments.

With respect to cell death induction, it is interesting to note that Cutler and

co-workers reported the accumulation of a GFP-Nitrilase 1 (GFP-Nit1) fusion protein in

Arabidopsis cells in response to mechanical wounding (Cutler and Somerville, 2005).

Plant nitrilases are reported to be involved in processes of cyanide detoxification, in

the catabolism of cyanogenic glycosides and glycosinolates, and in biosynthesis of the

plant growth hormone indole-3-acetic acid (Piotrowski, 2008). The appearance of

wound-induced vesiculation of GFP-Nit1 resembles strongly the accumulation of

14-3-3 dimers in punctate bodies observed in this study. In the co-localisation

analysis for 14-3-3 dimers and an ER marker presented here (Section 6.2.1), the

morphology of the ER marker and 14-3-3 dimers in epidermal peel cells was often

observed in vesicular structures. By contrast in intact leaves, it commonly presented

as a reticulated network of ER (Figs. 6-4 and 6-5).

The aggregation of GFP-Nit1 occurs rapidly from initially dispersed cytosolic

distribution upon wounding and cell death inductive treatments (Cutler and

Somerville, 2005). Thus, not just the final appearance of GFP-Nit1 aggregates but

also the timing upon wounding leading to such aggregates resembles strongly the

observations made for 14-3-3 dimers. The aggregation of GFP-Nit1 was proposed to


be an early marker of cell death at wound sites (Cutler and Somerville, 2005). More

interestingly still, a 14-3-3 interactome study has shown that Nit1 is a potential 14-3-3

target protein (Paul et al., 2009). Ongoing work in the Martin lab has demonstrated

interaction of several 14-3-3 isoforms with Nit1 in the cytosol and that the Nit1-14-3-3

complex aggregates into vesicular structures after wounding (M. van der Kwast and R.

Li, unpublished results). This suggests regulation of Nit1 by 14-3-3 proteins. It

further promotes the notion that 14-3-3s are somewhat involved in responses to cell

death triggers in plant cells. Whether such involvement is regulatory, i.e. impacts on

the progression of cell death events, or is simply a regulation of enzymes involved in

scavenging cellular components, e.g. recycling of nitrogen and detoxification of

cyanide compounds as performed by nitrilase, is unclear.

Chapter 7. General discussion 183

Chapter 7

General Discussion


7.1. Conclusion and discussion

7.1.1. Analysis of 14-3-3 isoform specificity using bimolecular fluorescence

complementation

Using the BiFC assay systems developed in the present study, dimerisation of

Arabidopsis 14-3-3 isoforms and binding of 14-3-3 proteins to the target protein

histone deacetylase 2C (HD2C) were investigated and visualised in living plant cells.

The work performed here showed that BiFC assays can be successfully performed in

transiently transformed Arabidopsis mesophyll protoplasts (Fig. 3-9),

Agrobacterium-infiltrated Nicotiana leaves (Fig. 3-10 to 3-13 and Chapter 4) and in

stably transformed Arabidopsis plants (Chapter 5).

The question whether the large number of 14-3-3 isoforms in multicellular

organisms reflects functional specificity (Rosenquist et al., 2000) was addressed in this

project by investigating: (1) if 14-3-3 proteins are selective for each other during dimer

formation and (2) if 14-3-3 isoform specificity exists in the interaction with the target

protein HD2C. In addition, the question was asked whether or not individual dimer

combinations have specific subcellular localisations, which could be an indicator of

functional specificity.

7.1.1.1. Isoform specificity in 14-3-3 dimerisation

Selectivity of a 14-3-3 isoform for another isoform during dimer formation could

be one of the mechanisms which contribute to 14-3-3 specificity (Aitken, 2002). To

test this hypothesis, ten Arabidopsis 14-3-3 isoforms were systematically examined for

their ability to dimerise with each other using transient BiFC assays in Nicotiana leaves

(Chapter 4). The results indicated that all ten 14-3-3 isoforms can freely dimerise

with each other (Chapter 4), suggesting little selectivity of Arabidopsis 14-3-3s during

dimer formation. This is in agreement with a number of previous studies


demonstrating for example that six Arabidopsis 14-3-3 isoforms can dimerise freely

with each other when using in vitro dimerisation assays or yeast two-hybrid studies

(Wu et al., 1997b; Abarca et al., 1999). The results of the present study also agree

with a recently published study showing that Arabidopsis 14-3-3 omega is able to

associate with ten Arabidopsis 14-3-3 isoforms in tandem affinity purification

experiments (Chang et al., 2009). Such results imply that formation of 14-3-3 dimers

contributes little to functional specificity of Arabidopsis 14-3-3 proteins. However,

this is in contrast to observations made in mammalian and yeast systems (Chaudhri et

al., 2003; Wilker et al., 2005). For example, the mammalian sigma isoform shows

selectivity, appearing to only form homodimers (Wilker et al., 2005). The

discrepancy to the results obtained here simply may be due to plant 14-3-3s behaving

differently to mammalian and yeast isoforms during dimer formation.

7.1.1.2. Isoform specificity with respect to 14-3-3 isoform-target interaction

Specificity of 14-3-3 proteins could be expressed by selectivity in the interaction

with target proteins (Rosenquist et al., 2000; Aitken, 2002). Differential binding

affinity between 14-3-3 isoforms and target proteins/peptides was shown using in

vitro binding assays and yeast two-hybrid analyses (Bachmann et al., 1996b;

Kanamaru et al., 1999; Rosenquist et al., 2000; Sullivan et al., 2009). In the present

study, interactions of seven Arabidopsis 14-3-3 isoforms with HD2C were investigated

for isoform specificity using transient BiFC analysis. All seven 14-3-3 isoforms tested

were able to interact with HD2C, implying a lack of isoform specificity. Similar results

have been obtained in our lab when testing for interaction of 14-3-3s with other

target proteins (R. Li, M. van der Kwast and T. Martin, unpublished results). One

possible explanation for the apparent lack of specificity may be that CaMV 35S

promoter-mediated high expression levels of 14-3-3 and target protein negatively

impact on selectivity. It is also possible that highly conserved ligand-binding


domains of 14-3-3 proteins cause low intrinsic differences in their ability to bind to

some ligands but may be sufficient for selectivity towards other ligands (Aitken et al.,

2002). The latter argument is supported by findings that 14-3-3 proteins show

specificity in the interaction with target proteins. For example, a large scale yeast

two-hybrid analysis testing five barley 14-3-3 isoforms for interaction with 132

potential 14-3-3 target proteins from barley indicated that some targets showed high

affinity with all five 14-3-3 isoforms while others interacted only with subsets of

14-3-3 isofomrs (Schoonheim et al., 2007a).

The internal conditions of the test system employed may also have impacted on

the selectivity of 14-3-3 isoforms towards a target protein. An example of this is the

ability of 14-3-3s to interact with the PM H+-ATPase (Alsterfjord et al., 2004).

Despite of the fact that twelve Arabidopsis 14-3-3 isoforms were able to bind to the

H+-ATPase in vitro, only a subset of the isoforms were found to associate with the

H+-ATPase in vivo (Alsterfjord et al., 2004). However, when a fusicoccin treatment

was applied to change the internal conditions, only 14-3-3 phi and upsilon isoforms,

which were not detected in the plasma membrane fraction under normal conditions,

were found to interact with the H+-ATPase (Alsterfjord et al., 2004). Thus,

interaction specificity of 14-3-3 isoforms may be target protein- or stimuli-dependent

and needs to be verified empirically case by case.

7.1.1.3. Differential subcellular distribution of 14-3-3 dimers may

contribute to specificity

In transient BiFC assays all 14-3-3 dimers were found in the cytoplasm and most

were also present in the nucleus of Nicotiana leave cells. Whilst the fluorescence

intensity for one dimer and between dimers in the cytoplasm of individual cells was

relatively constant, the nuclear fluorescence intensity varied. This was used to

classify cells expressing 14-3-3 dimers according to the nuclear to cytoplasmic BiFC


fluorescence ratios with three main patterns identified: Pattern 1 where

fluorescence in the nucleus was at least 1.5 times stronger than in the cytoplasm,

Pattern 2 where the ratio of fluorescence was between 1.5 and 0.7, and Pattern 3

with nuclear fluorescence far lower (ratio smaller than 0.7) than in the cytoplasm.

Most dimers appeared in more than one of these patterns, with one of the patterns

being pre-dominant. For example, dimers consisting of the kappa isoform and any

of the 14-3-3 isoforms epsilon, kappa, mu, omicron, phi and psi, were mostly

detected in Pattern 1. However, when kappa was paired with either iota, chi, nu or

omega, the dimers appeared preferentially in Pattern 2. In contrast to kappa

containing dimers, 14-3-3 nu-related dimers were usually observed in Pattern 2 or 3.

One interpretation of this is that preferential localisation may reflect a major role of

14-3-3 kappa in the regulation of nuclear target proteins while 14-3-3 nu may be

involved largely in the regulation of cytoplasmic target proteins.

When examining subcellular distribution of epsilon-YN/mu-YC, mu-YC/chi-YN

and mu-YN/mu-YC in transgenic Arabidopsis plants, dimer-dependent localisation

patterns were observed in some tissues. For example, the epsilon-YN/mu-YC

heterodimer was found in the nuclei of rosette leaf epidermal cells, root cells and

cotyledon guard cells whilst the mu-YN/mu-YC homodimer was not detectable in

nuclei of these three cell types (Figs. 5-5 to 5-7). This supported the hypothesis that

nuclear localisation is at least in part dimer-dependent.

A discrepancy between experiments performed in Nicotiana and Arabidopsis

was observed for mu homodimers, which may point to cell type, plant or

experimental system specific differences. For example, in contrast to the mu

homodimer, which was found in leaf cell nuclei during transient Nicotiana

experiments (Fig. 4-1), it was absent from nuclei of Arabidopsis rosette leaf cells (Fig.

5-7, J-L).


In summary, although this study found no evidence for specificity of 14-3-3

monomers for other monomers, during dimer formation, a level of specificity was

observed when subcellular localisation of 14-3-3 dimers was examined. This

specificity appeared not to be solely dependent on the dimer but may be strongly

influenced by cellular conditions. Thus, the physiological history of the cell and its

state at the time of experimentation may have a stronger impact on mediating

specificity of 14-3-3 dimers than the actual 14-3-3 protein itself.

7.1.2. Advantages and limitations of bimolecular fluorescence

complementation for the analysis of 14-3-3 interactions

Advantages and disadvantages of BiFC approaches have been discussed in the

literature (Walter et al., 2004; Kerppola, 2006; Ohad et al., 2007; Citovsky et al., 2008;

Kerppola, 2008; Lalonde et al., 2008). Briefly, the advantages of the BiFC assay are:

(1) It is an easy assay to perform that does not require sophisticated equipment for

the analysis of protein-protein interactions in vivo. (2) It allows live cell imaging of

interacting protein complexes, which prevents potential artefacts associated with cell

lysis or fixation. In addition, the subcellular localisation of such complexes can be

determined. (3) The assay is sensitive and capable of trapping weak or transient

interactions by the stable reconstitution of YFP complexes. (4) It has the potential to

detect the effects of stimuli on protein interactions.

The current study supports the reported advantages of BiFC assays. For

example, the dimerisations between 14-3-3 isoforms and the interactions of 14-3-3

isoforms with a target protein HD2C were easy to detect and identification of

subcellular localisations was achieved in living plant cells (Chapter 4).

On the other hand, limitations of BiFC assay may include: (1) The differences in

protein expression levels under a set of given experimental conditions probably


influence which binding partners can be identified (Kerppola, 2006). This can be

partially overcome by using strong promoters, although gene copy number still

impacts on expression. (2) There is a time delay between injection and formation of

the BiFC complex due to the need for transcription, translation and potential

pot-translational modifications to take place, in addition to time required for the

fluorophore to mature (Hu et al., 2002; Ohad et al., 2007). Thus, proteins with high

turnover rates or with large differences in stability might not be amenable to BiFC

studies (Bhat et al., 2006). (3) The formation of BiFC complexes is irreversible, at

least in vitro (Hu et al., 2002; Magliery et al., 2005). This characteristic might lead to

artificial interactions and hence false positive results, especially when overexpressing

such fusion proteins (Bhat et al., 2006; Lalonde et al., 2008). On the other hand the

tendency of the fluorescent protein fragments to associate is often reduced when

they are fused to proteins that do not associate with each other (Kerppola, 2008).

Thus, false positive interaction can be circumvented by performing appropriate

negative controls, for example using mutated proteins that are unable to interact.

The applicability of this for 14-3-3s was described in the literature (Walter et al., 2004;

Kerppola, 2008) and also in this thesis. Alternatively, the use of moderate promoters

for the expression of the respective fusion proteins will reduce such artefacts (Walter

et al., 2004; Bhat et al., 2006). (4) Steric accessibility of the split YFP parts in the

respective fusion proteins for reconstitution is an issue for the formation of BiFC

complexes (Lalonde et al., 2008). Thus, multiple fusion orientations should be tested

in case of negative outcomes (Morsy et al., 2008).

Most of the described limitations of BiFC assays did not impact on this study as

all investigated interactions occurred and were detectable whilst negative controls

with truncated, non-dimerising 14-3-3 mutants showed no interactions (Chapter 3).

However, some other potentially limiting factors need to be considered. Firstly, the


infiltration of Nicotiana leaves with a mixture of distinct Agrobacterium strains (YN, YC

and p19) restricted the use of the assay to qualitative examination of 14-3-3

interactions. This is due to the unpredictability of how the individual constructs will

be transferred and expressed in individual cells. Western blot analysis can only give

an averaged picture of protein expression in a tissue and does not show the

cell-to-cell differences. In theory, interactions can be analysed quantitatively by

directly comparing BiFC fluorescence intensity. This may be best achieved by

construction of a vector containing both, the YN and the YC fusion construct, thereby

reducing variability in transformations of two constructs. On the other hand, such

large vectors would be harder to handle and also greatly reduce the flexibility of the

approach, i.e. the combinatorial aspect, unless an increase in labour time for

generating more constructs is not an issue. Alternatively, adding an additional

reporter gene such as luciferase into the BiFC vectors would allow correction for

transformation differences, and may be an easier alternative to the above.

Secondly, it is unclear if the affinity of two interaction partners corresponds to

the degree of cellular fluorescence (Bhat et al., 2006). Thus observed BiFC

fluorescence of different 14-3-3 interactions cannot be directly used as an indicator

for the interaction preference between distinct 14-3-3 dimeric combinations or

different 14-3-3 isoform-target interactions. Although the intensity of BiFC

fluorescence per se is quantifiable (Walter et al., 2004; Gampala et al., 2007), it

requires an internal control to compare the intensity between cells. The multicolour

BiFC analysis, which allows simultaneous visualisation of multiple protein complexes

in the same cell (Hu and Kerppola, 2003; Kerppola, 2008), has the potential to address

competition between two or more 14-3-3 proteins for a target protein which may be

another 14-3-3 or a target protein. However, the multicolour BiFC analysis is still a

challenging technique as it requires precise quantification of the amount of


fluorescent protein complexes with often overlapping excitation and/or emission

spectra and big differences in fluorescence energy output hindering visualisation and

signal comparision.

Last but not least, it may not be possible to resolve the full function of 14-3-3

proteins, especially in terms of dissociation of the protein complexes, using the BiFC

assays because the formation of BiFC complexes is irreversible (Hu et al., 2002;

Magliery et al., 2005). On the other hand, it is speculated that the dynamics of BiFC

complex formation is maintained or can be modified by the cellular protein folding

and degradation machinery, allowing at least the stability versus degradation of BiFC

complexes to be investigated (Walter et al., 2004). Furthermore, intracellular

relocalisation, such as those observed in this project in response to wounding and cell

death (Chapter 4 and 6), is possible with this system.

7.1.3. Interpretation of the BiFC outcomes of 14-3-3 interactions in cells

This study showed the ability of the introduced 14-3-3-split YFP fusion proteins

to bind to their interacting counterparts in the developed BiFC assay systems. Thus,

it can be speculated that an introduced 14-3-3 fusion protein is also able to interact

with endogenous 14-3-3 interacting proteins, including native 14-3-3 isoforms and

their target proteins. From a different point of view, the introduced 14-3-3s are in

direct competition with endogenous 14-3-3s for dimerisation and interaction with

targets.

In the transient BiFC assay of Nicotiana leaves, it can be expected that the

amounts of expressed 14-3-3-split YFP fusion molecules may be much higher than

those of the endogenous 14-3-3 interacting proteins, owing to induction of the

transgene expression by the co-expression of silencing suppressor protein p19. Thus,

intrinsic competition may be low and the BiFC signals would largely reflect the two


interacting proteins been tested. This may also explain the contradictory BiFC

distribution patterns that showed all the examined 14-3-3 dimerisations were absent

from the nucleolus whilst 14-3-3-HD2C interactions accumulated to high levels in the

nucleolus (Fig. 4-7). The absence of 14-3-3 dimers from nucleoli in dimerisation

experiments may be a mere artefact based on the lack of a similarly highly expressed

target protein. Once a target protein is available in high numbers, it may provide the

driving force, bringing 14-3-3s into the nucleolus. Complementing dimerisation

studies with localisation studies using single 14-3-3 isoforms fused to GFP or another

marker protein may help avoid mis- or over-interpretation of dimerisation data. This

approach may be limited to some isoforms as it was reported that out of six isoforms,

only kappa localised to the nucleolus when using a GFP fusion protein approach

(Koroleva et al., 2005; Paul et al., 2005; Rienties et al., 2005; Reumann et al., 2009;

Table 4-1).

In contrast, for the transgenic Arabidopsis plants used with the BiFC assay, the

expression level of transgenes could be different from plant to plant due to the

balance between the effects of transgene-driven promoter and the intrinsic gene

expression regulatory mechanisms such as post transcriptional gene silencing

(Wroblewski et al., 2005). Besides, a reservoir of endogenous 14-3-3 interacting

proteins may act as competitors for, or may disrupt the formation of, recombinant

fluorescent BiFC complexes in cells (MacKintosh, 2004). Thus, if there are any

preferential dimerisations between 14-3-3 isoforms or isoform-specific interactions of

14-3-3 with target proteins, it would have a negative impact on formation of the

tested 14-3-3 BiFC complexes in a particular cellular compartment in the native

system. This may explain why the mu-YN/mu-YC dimer (found in the nuclei in the

Nicotiana system) was not detectable in the nucleus of leaf epidermal cells and

cotyledon guard cells in the transgenic Arabidopsis plants but the mu-YC/chi-YN and


epsilon-YN/mu-YC heterodimers were (Figs. 5-6 and 5-7). Hence it would be

important to know the protein levels of the transgenic 14-3-3-split YFP over-expressed

related to the endogenous 14-3-3 isoforms. For this, proper tools, such as 14-3-3

isoform-specific antibodies, are needed.

7.2. Future perspectives

Despite hundreds of 14-3-3 target proteins having been predicted, their

interaction with 14-3-3s in vivo or in the native organism is still speculative (Oecking

and Jaspert, 2009). As was shown elsewhere (Walter et al., 2004; Gampala et al.,

2007; Jaspert et al., 2009; Purwestri et al., 2009), BiFC is a suitable method to provide

in vivo or in planta evidence for such interactions. Isoform specificity of 14-3-3

proteins was not observed in the present study, neither in the dimerisation

experiments nor in the interaction studies with HD2C. It is possible that isoforms

omitted here would reveal specificity when tested for dimerisation or HD2C

interaction, and these should be tested.

To ensure that the observed interactions with HD2C are not due to non-specific

binding, negative controls such as N-terminally truncated 14-3-3s could be used. A

further control would be to use a HD2C mutant in which the 14-3-3 binding site was

mutated. The first set of experiments was conducted in the Martin lab and no

interaction was detected (M. van der Kwast and T. Martin, unpublished data). With

respect to the second approach, deletion experiments were used to show that the

potential 14-3-3 binding site is located in the N-terminal region of the HD2C (M. van

der Kwast and T. Martin, unpublished data).

If the 14-3-3 BiFC dimeric complex is still able to interact with target proteins, i.e.

the YFP components do not prevent such interaction, transgenic plants could be used

along with pull-down assays in combination with two-dimensional gel electrophoresis


and gas chromatography/mass spectrometry (GC/MS) or MS alone to identify target

proteins binding to the 14-3-3-split YFP fusion proteins. The library of transgenic

Arabidopsis plants carrying one or two 14-3-3 BiFC constructs can be used as

biological starting material to generate such 14-3-3 interactomic profiles for different

tissues or different development stages. Similar strategies of using antibodies to

pull-down 14-3-3 associated protein complexes have been successfully performed

(Chang et al., 2009; Paul et al., 2009). The association of -YN and -YC constructs with

two different tags may be suitable to identify target proteins interacting with the

dimer only but not with single 14-3-3s or one of the 14-3-3 BiFC constructs and an

endogenous 14-3-3 protein. This, in principle, allows for a proteomics approach

directed towards the identification of dimer specific binding partners.

Alternatively, an approach that can isolate intact protein complexes, such as

blue-native gel electrophoresis (Kanczewska et al., 2005; Wittig et al., 2006) could be

used as the first step in the identification of components of 14-3-3-interacting protein

complex.

Changes in cellular distribution of 14-3-3 BiFC fluorescence can occur in

response to certain cellular events as was demonstrated in this work when studying

14-3-3 dimer distribution in response to cell death-related stimuli (Chapter 4 and 6).

This has since led to the discovery that 14-3-3-Nit1 interactions were commonly in the

cytoplasm, but aggregation of the 14-3-3-Nit1 BiFC complex into vesicular structures

was induced by wounding and herbicide treatments (R. Li, M. van der Kwast and T.

Martin, unpublished results). Future studies could also include examining the links

between 14-3-3 interactions with other target proteins, changes in subcellular

distribution, and the biological role of 14-3-3s or their target proteins.

In conclusion, BiFC assays adapted in this thesis were successfully used to

investigate the in vivo interactions and subcellular localisations of 14-3-3 proteins in


dimerisation studies and with a target protein. It was found that this technique has

great potential to investigate 14-3-3-target complex formation and translocation in

living cells. The present study also led to investigations in the potential roles of

14-3-3 proteins in regulation of nitrilase and HD2C. Further studies in these

directions will provide novel insights into the complex roles of 14-3-3 proteins.

References 196

References

Abarca, D., Madueno, F., Martinez-Zapater, J.M., and Salinas, J. (1999). Dimerization

of Arabidopsis 14-3-3 proteins: Structural requirements within the N-terminal

domain and effect of calcium. FEBS Lett. 462, 377-382.

Aitken, A. (1996). 14-3-3 and its possible role in co-ordinating multiple signalling

pathways. Trends Cell Biol. 6, 341-347.

Aitken, A. (2002). Functional specificity in 14-3-3 isoform interactions through dimer

formation and phosphorylation. Chromosome location of mammalian isoforms

and variants. Plant Mol. Biol. 50, 993-1010.

Aitken, A., Baxter, H., Dubois, T., Clokie, S., Mackie, S., Mitchell, K., Peden, A., and

Zemlickova, E. (2002). Specificity of 14-3-3 isoform dimer interactions and

phosphorylation. Biochem. Soc. Trans. 30, 351-360.

Alsterfjord, M., Sehnke, P.C., Arkell, A., Larsson, H., Svennelid, F., Rosenquist, M.,

Ferl, R.J., Sommarin, M., and Larsson, C. (2004). Plasma membrane H+-ATPase

and 14-3-3 isoforms of Arabidopsis leaves: Evidence for isoform specificity in

the 14-3-3/H+-ATPase interaction. Plant Cell Physiol. 45, 1202-1210.

Aparicio, F., Thomas, C.L., Lederer, C., Niu, Y., Wang, D., and Maule, A.J. (2005). Virus

induction of heat shock protein 70 reflects a general response to protein

accumulation in the plant cytosol. Plant Physiol. 138, 529-536.

Asai, T., Stone, J.M., Heard, J.E., Kovtun, Y., Yorgey, P., Sheen, J., and Ausubel, F.M.

(2000). Fumonisin B1-induced cell death in Arabidopsis protoplasts requires

jasmonate-, ethylene-, and salicylate-dependent signaling pathways. Plant Cell

12, 1823-1836.

Athwal, G.S., and Huber, S.C. (2002). Divalent cations and polyamines bind to loop 8

of 14-3-3 proteins, modulating their interaction with phosphorylated nitrate

reductase. Plant J. 29, 119-129.

Athwal, G.S., Huber, J.L., and Huber, S.C. (1998a). Biological significance of divalent

metal ion binding to 14-3-3 proteins in relationship to nitrate reductase

inactivation. Plant Cell Physiol. 39, 1065-1072.

Athwal, G.S., Huber, J.L., and Huber, S.C. (1998b). Phosphorylated nitrate reductase

and 14-3-3 proteins. Site of interaction, effects of ions, and evidence for an

AMP-binding site on 14-3-3 proteins. Plant Physiol. 118, 1041-1048.

Athwal, G.S., Lombardo, C.R., Huber, J.L., Masters, S.C., Fu, H., and Huber, S.C. (2000).

Modulation of 14-3-3 protein interactions with target polypeptides by physical

and metabolic effectors. Plant Cell Physiol. 41, 523-533.

Bachmann, M., Huber, J.L., Liao, P.-C., Gage, D.A., and Huber, S.C. (1996a). The

inhibitor protein of phosphorylated nitrate reductase from spinach (Spinacia

oleracea) leaves is a 14-3-3 protein. FEBS Lett. 387, 127-131.

References 197

Bachmann, M., Huber, J.L., Athwal, G.S., Wu, K., Ferl, R.J., and Huber, S.C. (1996b).

14-3-3 proteins associate with the regulatory phosphorylation site of spinach

leaf nitrate reductase in an isoform-specific manner and reduce

dephosphorylation of Ser-543 by endogenous protein phosphatases. FEBS Lett.

398, 26-30.

Bae, M.S., Cho, E.J., Choi, E.-Y., and Park, O.K. (2003). Analysis of the Arabidopsis

nuclear proteome and its response to cold stress. Plant J. 36, 652-663.

Baunsgaard, L., Fuglsang, A.T., Jahn, T., Korthout, H.A.A.J., de Boer, A.H., and

Palmgren, M.G. (1998). The 14-3-3 proteins associate with the plant plasma

membrane H+-ATPase to generate fusicoccin binding complex and a fusicoccin

responsive system. Plant J. 13, 661-671.

Beers, E.P., and McDowell, J.M. (2001). Regulation and execution of programmed cell

death in response to pathogens, stress and developmental cues. Curr. Opin.

Plant Biol. 4, 561-567.

Benschop, J.J., Mohammed, S., O'Flaherty, M., Heck, A.J.R., Slijper, M., and Menke,

F.L.H. (2007). Quantitative phosphoproteomics of early elicitor signaling in

Arabidopsis. Mol. Cell. Proteomics 6, 1198-1214.

Bhat, R.A., Lahaye, T., and Panstruga, R. (2006). The visible touch: in planta

visualization of protein-protein interactions by fluorophore-based methods.

Plant Methods 2, 12.

Bihn, E.A., Paul, A.-L., Wang, S.W., Erdos, G.W., and Ferl, R.J. (1997). Localization of

14-3-3 proteins in the nuclei of arabidopsis and maize. Plant J. 12, 1439-1445.

Bracha-Drori, K., Shichrur, K., Katz, A., Oliva, M., Angelovici, R., Yalovsky, S., and

Ohad, N. (2004). Detection of protein-protein interactions in plants using

bimolecular fluorescence complementation. Plant J. 40, 419-427.

Brandt, J., Thordal-Christensen, H., Vad, K., Gregersen, P.L., and Collinge, D.B. (1992).

A pathogen-induced gene of barley encodes a protein showing high similarity

to a protein kinase regulator. Plant J. 2, 815-820.

Braselmann, S., and McCormick, F. (1995). Bcr and Raf form a complex in vivo via

14-3-3 proteins. EMBO J. 14, 4839-4848.

Bridges, D., and Moorhead, G.B.G. (2005). 14-3-3 proteins: a number of functions for

a numbered protein. Sci. STKE 2005, re10.

Budnik, A., and Stephens, D.J. (2009). ER exit sites - Localization and control of COPII

vesicle formation. FEBS Lett. 583, 3796-3803.

Bunney, T.D., van Walraven, H.S., and de Boer, A.H. (2001). 14-3-3 Protein is a

regulator of the mitochondrial and chloroplast ATP synthase. Proc. Natl. Acad.

Sci. USA 98, 4249-4254.

Bunney, T.D., van den Wijngaard, P.W.J., and de Boer, A.H. (2002). 14-3-3 Protein

regulation of proton pumps and ion channels. Plant Mol. Biol. 50, 1041-1051.

References 198

Bustos, D.M., and Iglesias, A.A. (2003). Phosphorylated non-phosphorylating

glyceraldehyde-3-phosphate dehydrogenase from heterotrophic cells of wheat

interacts with 14-3-3 proteins. Plant Physiol. 133, 2081-2088.

Campbell, W.H. (1999). NITRATE REDUCTASE STRUCTURE, FUNCTION AND

REGULATION: bridging the gap between biochemistry and physiology. Annu.

Rev. Plant Physiol. Plant Mol. Biol. 50, 277-303.

Cao, A., Jain, A., Baldwin, J., and Raghothama, K. (2007). Phosphate differentially

regulates 14-3-3 family members and GRF9 plays a role in Pi-starvation

induced responses. Planta 226, 1219-1230.

Carbon, S., Ireland, A., Mungall, C.J., Shu, S., Marshall, B., Lewis, S., the Ami, G.O.H.,

and the Web Presence Working, G. (2009). AmiGO: online access to ontology

and annotation data. Bioinformatics 25, 288-289.

Chang, I.-F., Curran, A., Woolsey, R., Quilici, D., Cushman, J.C., Mittler, R., Harmon, A.,

and Harper, J.F. (2009). Proteomic profiling of tandem affinity purified 14-3-3

protein complexes in Arabidopsis thaliana. Proteomics 9, 2967-2985.

Chaudhri, M., Scarabel, M., and Aitken, A. (2003). Mammalian and yeast 14-3-3

isoforms form distinct patterns of dimers in vivo. Biochem. Biophys. Res.

Commun. 300, 679-685.

Chen, F., Li, Q., Sun, L., and He, Z. (2006). The rice 14-3-3 gene family and its

involvement in responses to biotic and abiotic stress. DNA Res 13, 53-63.

Chevalier, D., Morris, E.R., and Walker, J.C. (2009). 14-3-3 and FHA domains mediate

phosphoprotein interactions. Annu. Rev. Plant Biol. 60, 67-91.

Chung, H.-J., Sehnke, P.C., and Ferl, R.J. (1999). The 14-3-3 proteins: cellular

regulators of plant metabolism. Trends Plant Sci. 4, 367-371.

Citovsky, V., Gafni, Y., and Tzfira, T. (2008). Localizing protein-protein interactions by

bimolecular fluorescence complementation in planta. Methods 45, 196-206.

Clough, S.J., and Bent, A.F. (1998). Floral dip: a simplified method for

Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J. 16,

735-743.

Cohen, P. (2002). The origins of protein phosphorylation. Nat. Cell Biol. 4, E127-E130.

Comparot, S., Lingiah, G., and Martin, T. (2003). Function and specificity of 14-3-3

proteins in the regulation of carbohydrate and nitrogen metabolism. J. Exp. Bot.

54, 595-604.

Cutler, S., and Somerville, C. (2005). Imaging plant cell death: GFP-Nit1 aggregation

marks an early step of wound and herbicide induced cell death. BMC Plant Biol.

5, 4.

Cutler, S.R., Ehrhardt, D.W., Griffitts, J.S., and Somerville, C.R. (2000). Random

GFP::cDNA fusions enable visualization of subcellular structures in cells of

Arabidopsis at a high frequency. Proc. Natl. Acad. Sci. USA 97, 3718-3723.

References 199

Daugherty, C.J., Rooney, M.F., Miller, P.W., and Ferl, R.J. (1996). Molecular

organization and tissue-specific expression of an Arabidopsis 14-3-3 gene.

Plant Cell 8, 1239-1248.

de Boer, B. (1997). Fusicoccin - a key to multiple 14-3-3 locks? Trends Plant Sci. 2,

60-66.

de Vetten, N.C., Lu, G., and Ferl, R.J. (1992). A maize protein associated with the

G-box binding complex has homology to brain regulatory proteins. Plant Cell 4,

1295-1307.

Desikan, R., Reynolds, A., Hancock, J.T., and Neill, S.J. (1998). Harpin and hydrogen

peroxide both initiate programmed cell death but have differential effects on

defence gene expression in Arabidopsis suspension cultures. Biochem. J. 330,

115-120.

Desprez, T., Juraniec, M., Crowell, E.F., Jouy, H., Pochylova, Z., Parcy, F., Höfte, H.,

Gonneau, M., and Vernhettes, S. (2007). Organization of cellulose synthase

complexes involved in primary cell wall synthesis in Arabidopsis thaliana. Proc.

Natl. Acad. Sci. USA 104, 15572-15577.

Dougherty, M.K., and Morrison, D.K. (2004). Unlocking the code of 14-3-3. J. Cell Sci.

117, 1875-1884.

Douglas, P., Moorhead, G., Hong, Y., Morrice, N., and MacKintosh, C. (1998).

Purification of a nitrate reductase kinase from Spinacea oleracea leaves, and

its identification as a calmodulin-domain protein kinase. Planta 206, 435-442.

Eckardt, N.A. (2001). Transcription factors dial 14-3-3 for nuclear shuttle. Plant Cell 13,

2385-2389.

Emanuelsson, O., Elofsson, A., von Heijne, G., and Cristobal, S. (2003). In silico

prediction of the peroxisomal proteome in fungi, plants and animals. J. Mol.

Biol. 330, 443-456.

Ferl, R.J. (2004). 14-3-3 proteins: Regulation of signal-induced events. Physiol. Plant.

120, 173-178.

Ferl, R.J., Manak, M.S., and Reyes, M.F. (2002). The 14-3-3s. Genome Biol. 3,

reviews3010.

Finnemann, J., and Schjoerring, J.K. (2000). Post-translational regulation of cytosolic

glutamine synthetase by reversible phosphorylation and 14-3-3 protein

interaction. Plant J. 24, 171-181.

Fu, H., Subramanian, R.R., and Masters, S.C. (2000). 14-3-3 proteins: structure,

function, and regulation. Annu. Rev. Pharmacol. Toxicol. 40, 617-647.

Gampala, S.S., Kim, T.-W., He, J.-X., Tang, W., Deng, Z., Bai, M.-Y., Guan, S., Lalonde,

S., Sun, Y., Gendron, J.M., Chen, H., Shibagaki, N., Ferl, R.J., Ehrhardt, D.,

Chong, K., Burlingame, A.L., and Wang, Z.-Y. (2007). An essential role for

14-3-3 proteins in brassinosteroid signal transduction in Arabidopsis. Dev. Cell

13, 177-189.

References 200

Gardino, A.K., Smerdon, S.J., and Yaffe, M.B. (2006). Structural determinants of

14-3-3 binding specificities and regulation of subcellular localization of

14-3-3-ligand complexes: A comparison of the X-ray crystal structures of all

human 14-3-3 isoforms. Semin. Cancer Biol. 16, 173-182.

Gawel, N.J., and Jarret, R.L. (1991). A modified CTAB DNA extraction procedure for

Musa and Ipomoea. Plant Mol. Biol. Report. 9, 262-266.

Genoud, T., and Métraux, J.-P. (1999). Crosstalk in plant cell signaling: structure and

function of the genetic network. Trends Plant Sci. 4, 503-507.

Gomord, V., Denmat, L.-A., Fitchette-Lainé, A.-C., Satiat-Jeunemaitre, B., Hawes, C.,

and Faye, L. (1997). The C-terminal HDEL sequence is sufficient for retention of

secretory proteins in the endoplasmic reticulum (ER) but promotes vacuolar

targeting of proteins that escape the ER. Plant J. 11, 313-325.

Grefen, C., Stadele, K., Ruzicka, K., Obrdlik, P., Harter, K., and Horak, J. (2008).

Subcellular localization and in vivo interactions of the Arabidopsis thaliana

ethylene receptor family members. Mol Plant 1, 308-320.

Grozinger, C.M., and Schreiber, S.L. (2000). Regulation of histone deacetylase 4 and 5

and transcriptional activity by 14-3-3-dependent cellular localization. Proc.

Natl. Acad. Sci. USA 97, 7835-7840.

Höfgen, R., and Willmitzer, L. (1988). Storage of competent cells for Agrobacterium

transformation. Nucl. Acids Res. 16, 9877.

Hamilton, A., Voinnet, O., Chappell, L., and Baulcombe, D. (2002). Two classes of

short interfering RNA in RNA silencing. EMBO J. 21, 4671-4679.

Hanahan, D. (1985). Techniques for transformation of E. coli. In DNA Cloning: a

practical approach, D.M. Glover, ed (London, UK: IRL Press Ltd.

Hanton, S.L., Bortolotti, L.E., Renna, L., Stefano, G., and Brandizzi, F. (2005). Crossing

the divide - transport between the endoplasmic reticulum and Golgi apparatus

in plants. Traffic 6, 267-277.

Harrison, S.J., Mott, E.K., Parsley, K., Aspinall, S., Gray, J.C., and Cottage, A. (2006). A

rapid and robust method of identifying transformed Arabidopsis thaliana

seedlings following floral dip transformation. Plant Methods 2, 19.

He, Z.-H., Cheeseman, I., He, D., and Kohorn, B. (1999). A cluster of five cell

wall-associated receptor kinase genes, Wak1–5, are expressed in specific

organs of Arabidopsis. Plant Mol. Biol. 39, 1189-1196.

Heazlewood, J.L., Verboom, R.E., Tonti-Filippini, J., Small, I., and Millar, A.H. (2007).

SUBA: the Arabidopsis subcellular database. Nucl. Acids Res. 35, D213-D218.

Helenius, E., Boije, M., Niklander-Teeri, V., Palva, E., and Teeri, T. (2000). Gene

delivery into intact plants using the HeliosTM gene gun. Plant Mol. Biol. Report.

18, 287-288.

Hellens, R., Mullineaux, P., and Klee, H. (2000a). A guide to Agrobacterium binary Ti

vectors. Trends Plant Sci. 5, 446-451.

References 201

Hellens, R.P., Edwards, E.A., Leyland, N.R., Bean, S., and Mullineaux, P.M. (2000b).

pGreen: a versatile and flexible binary Ti vector for Agrobacterium-mediated

plant transformation. Plant Mol. Biol. 42, 819-832.

Hellens, R.P., Allan, A.C., Friel, E.N., Bolitho, K., Grafton, K., Templeton, M.D.,

Karunairetnam, S., Gleave, A.P., and Laing, W.A. (2005). Transient expression

vectors for functional genomics, quantification of promoter activity and RNA

silencing in plants. Plant Methods 1, 13.

Hirsch, S., Aitken, A., Bertsch, U., and Soll, J.g. (1992). A plant homologue to

mammalian brain 14-3-3 protein and protein kinase C inhibitor. FEBS Lett. 296,

222-224.

Hoff, B., and Kück, U. (2005). Use of bimolecular fluorescence complementation to

demonstrate transcription factor interaction in nuclei of living cells from the

filamentous fungus Acremonium chrysogenum. Curr. Genet. 47, 132-138.

Hollender, C., and Liu, Z. (2008). Histone deacetylase genes in Arabidopsis

development. J. Integr. Plant Biol. 50, 875-885.

Horton, P., Park, K.-J., Obayashi, T., Fujita, N., Harada, H., Adams-Collier, C.J., and

Nakai, K. (2007). WoLF PSORT: protein localization predictor. Nucl. Acids Res.

35, W585-587.

Houot, V., Etienne, P., Petitot, A.-S., Barbier, S., Blein, J.-P., and Suty, L. (2001).

Hydrogen peroxide induces programmed cell death features in cultured

tobacco BY-2 cells, in a dose-dependent manner. J. Exp. Bot. 52, 1721-1730.

Howden, R., Park, S.K., Moore, J.M., Orme, J., Grossniklaus, U., and Twell, D. (1998).

Selection of T-DNA-tagged male and female gametophytic mutants by

segregation distortion in Arabidopsis. Genetics 149, 621-631.

Hu, C.-D., and Kerppola, T.K. (2003). Simultaneous visualization of multiple protein

interactions in living cells using multicolor fluorescence complementation

analysis. Nat. Biotechnol. 21, 539-545.

Hu, C.-D., Chinenov, Y., and Kerppola, T.K. (2002). Visualization of interactions among

bZIP and Rel family proteins in living cells using bimolecular fluorescence

complementation. Mol. Cell 9, 789-798.

Hua, S., and Sun, Z. (2001). Support vector machine approach for protein subcellular

localization prediction. Bioinformatics 17, 721-728.

Huang, C.-N., Cornejo, M.J., Bush, D.S., and Jones, R.L. (1986). Estimating viability of

plant protoplasts using double and single staining. Protoplasma 135, 80-87.

Huang, H.C., and Chen, C.C. (1988). Genome Multiplication in Cultured Protoplasts of

Two Nicotiana Species. J. Hered. 79, 28-32.

Huber, S.C., Bachmann, M., and Huber, J.L. (1996). Post-translational regulation of

nitrate reductase activity: a role for Ca2+ and 14-3-3 proteins. Trends Plant Sci.

1, 432-438.

References 202

Huber, S.C., MacKintosh, C., and Kaiser, W.M. (2002). Metabolic enzymes as targets

for 14-3-3 proteins. Plant Mol. Biol. 50, 1053-1063.

Ichimura, T., Isobe, T., Okuyama, T., Yamauchi, T., and Fujisawa, H. (1987). Brain

14-3-3 protein is an activator protein that activates tryptophan

5-monooxygenase and tyrosine 3-monooxygenase in the presence of

Ca2+,calmodulin-dependent protein kinase II. FEBS Lett. 219, 79-82.

Igarashi, D., Ishida, S., Fukazawa, J., and Takahashi, Y. (2001). 14-3-3 Proteins

regulate intracellular localization of the bZIP transcriptional activator RSG.

Plant Cell 13, 2483-2497.

Jach, G., Pesch, M., Richter, K., Frings, S., and Uhrig, J.F. (2006). An improved mRFP1

adds red to bimolecular fluorescence complementation. Nat.Methods 3,

597-600.

Jahn, T., Fuglsang, A.T., Olsson, A., Bruntrup, I.M., Collinge, D.B., Volkmann, D.,

Sommarin, M., Palmgren, M.G., and Larsson, C. (1997). The 14-3-3 protein

interacts directly with the C-terminal region of the plant plasma membrane

H+-ATPase. Plant Cell 9, 1805-1814.

Jaquinod, M., Villiers, F., Kieffer-Jaquinod, S., Hugouvieux, V., Bruley, C., Garin, J.,

and Bourguignon, J. (2007). A proteomics dissection of Arabidopsis thaliana

vacuoles isolated from cell culture. Mol. Cell. Proteomics 6, 394-412.

Jaspert, N., and Oecking, C. (2002). Regulatory 14-3-3 proteins bind the atypical motif

within the C terminus of the plant plasma membrane H+-ATPase via their

typical amphipathic groove. Planta 216, 136-139.

Jaspert, N., Weckermann, K., Piotrowski, M., and Oecking, C. (2009). Enolase, a

cross-link between glycolysis and stress response. In 20th International

Conference on Arabidopsis Research (Edinburgh, Scotland, United Kingdom).

Jones, A.M. (2001). Programmed cell death in development and defense. Plant

Physiol. 125, 94-97.

Jones, D.H., Ley, S., and Aitken, A. (1995). Isoforms of 14-3-3 protein can form homo-

and heterodimers in vivo and in vitro: implications for function as adapter

proteins. FEBS Lett. 368, 55-58.

Köhler, R.H., Zipfel, W.R., Webb, W.W., and Hanson, M.R. (1997). The green

fluorescent protein as a marker to visualize plant mitochondria in vivo. Plant J.

11, 613-621.

Kanamaru, K., Wang, R., Su, W., and Crawford, N.M. (1999). Ser-534 in the hinge 1

region of Arabidopsis nitrate reductase is conditionally required for binding of

14-3-3 proteins and in vitro inhibition. J. Biol. Chem. 274, 4160-4165.

Kanczewska, J., Marco, S., Vandermeeren, C., Maudoux, O., Rigaud, J.-L., and Boutry,

M. (2005). Activation of the plant plasma membrane H+-ATPase by

phosphorylation and binding of 14-3-3 proteins converts a dimer into a

hexamer. Proc. Natl. Acad. Sci. USA 102, 11675-11680.

References 203

Karlova, R., Boeren, S., Russinova, E., Aker, J., Vervoort, J., and de Vries, S. (2006).

The Arabidopsis SOMATIC EMBRYOGENESIS RECEPTOR-LIKE KINASE1 protein

complex includes BRASSINOSTEROID-INSENSITIVE1. Plant Cell 18, 626-638.

Kerppola, T.K. (2006). Visualization of molecular interactions by fluorescence

complementation. Nat. Rev. Mol. Cell Biol. 7, 449-456.

Kerppola, T.K. (2008). Bimolecular fluorescence complementation (BiFC) analysis as a

probe of protein interactions in living cells. Annu. Rev. Biophys. 37, 465-487.

Kleffmann, T., Russenberger, D., von Zychlinski, A., Christopher, W., Sjölander, K.,

Gruissem, W., and Baginsky, S. (2004). The Arabidopsis thaliana chloroplast

proteome reveals pathway abundance and novel protein functions. Curr. Biol.

14, 354-362.

Koncz, C., and Schell, J. (1986). The promoter of TL-DNA gene 5 controls the

tissue-specific expression of chimaeric genes carried by a novel type of

Agrobacterium binary vector. Mol. Gen. Genet. 204, 383-396.

Koroleva, O.A., Tomlinson, M.L., Leader, D., Shaw, P., and Doonan, J.H. (2005).

High-throughput protein localization in Arabidopsis using

Agrobacterium-mediated transient expression of GFP-ORF fusions. Plant J. 41,

162-174.

Kovtun, Y., Chiu, W.-L., Tena, G., and Sheen, J. (2000). Functional analysis of oxidative

stress-activated mitogen-activated protein kinase cascade in plants. Proc. Natl.

Acad. Sci. USA 97, 2940-2945.

Kubista, M., Aakerman, B., and Norden, B. (1987). Characterization of interaction

between DNA and 4',6-diamidino-2-phenylindole by optical spectroscopy.

Biochemistry 26, 4545-4553.

Kulma, A., Villadsen, D., Campbell, D.G., Meek, S.E.M., Harthill, J.E., Nielsen, T.H.,

and MacKintosh, C. (2004). Phosphorylation and 14-3-3 binding of Arabidopsis

6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase. Plant J. 37, 654-667.

Kuromori, T., and Yamamoto, M. (2000). Members of the Arabidopsis 14-3-3 gene

family trans-complement two types of defects in fission yeast. Plant Sci. 158,

155-161.

Laemmli, U.K. (1970). Cleavage of Structural Proteins during the Assembly of the

Head of Bacteriophage T4. Nature 227, 680-685.

Lalonde, S., Ehrhardt, D.W., Loque, D., Chen, J., Rhee, S.Y., and Frommer, W.B. (2008).

Molecular and cellular approaches for the detection of protein-protein

interactions: latest techniques and current limitations. Plant J. 53, 610-635.

Lam, E. (2004). Controlled cell death, plant survival and development. Nat. Rev. Mol.

Cell Biol. 5, 305-315.

Lapointe, G., Luckevich, M.D., Cloutier, M., and Seguin, A. (2001). 14-3-3 gene family

in hybrid poplar and its involvement in tree defence against pathogens. J. Exp.

Bot. 52, 1331-1338.

References 204

Lawrence, R.J., Earley, K., Pontes, O., Silva, M., Chen, Z.J., Neves, N., Viegas, W., and

Pikaard, C.S. (2004). A concerted DNA methylation/histone methylation switch

regulates rRNA gene dosage control and nucleolar dominance. Mol. Cell 13,

599-609.

Lillo, C., Lea, U.S., Leydecker, M.T., and Meyer, C. (2003). Mutation of the regulatory

phosphorylation site of tobacco nitrate reductase results in constitutive

activation of the enzyme in vivo and nitrite accumulation. Plant J. 35, 566-573.

Lillo, C., Meyer, C., Lea, U.S., Provan, F., and Oltedal, S. (2004). Mechanism and

importance of post-translational regulation of nitrate reductase. J. Exp. Bot. 55,

1275-1282.

Liu, D., Bienkowska, J., Petosa, C., Collier, R.J., Fu, H., and Liddington, R. (1995).

Crystal structure of the zeta isoform of the 14-3-3 protein. Nature 376,

191-194.

Logemann, E., Birkenbihl, R.P., Ü lker, B., and Somssich, I.E. (2006). An improved

method for preparing Agrobacterium cells that simplifies the Arabidopsis

transformation protocol. Plant Methods 2, 16.

Lu, G., Sehnke, P.C., and Ferl, R.J. (1994). Phosphorylation and calcium binding

properties of an Arabidopsis GF14 brain protein homolog. Plant Cell 6,

501-510.

Lu, G., DeLisle, A.J., de Vetten, N.C., and Ferl, R.J. (1992). Brain proteins in plants: an

Arabidopsis homolog to neurotransmitter pathway activators is part of a DNA

binding complex. Proc. Natl. Acad. Sci. USA 89, 11490-11494.

Lu, Z.-X., Laroche, A., and Huang, H.C. (2004). Segregation patterns for integration

and expression of Coniothyrium minitans xylanase gene in Arabidopsis thaliana

transformants. Bot. Bull. Acad. Sin. (Taipei) 45, 23-31.

Luo, Z.-j., Zhang, X.-f., Rapp, U., and Avruch, J. (1995). Identification of the 14.3.3 ζ

domains important for self-association and Raf binding. J. Biol. Chem. 270,

23681-23687.

MacKintosh, C. (2004). Dynamic interactions between 14-3-3 proteins and

phosphoproteins regulate diverse cellular processes. Biochem. J. 381, 329-342.

Mackintosh, C., Douglas, P., and Lillo, C. (1995). Identification of a protein that

inhibits the phosphorylated form of nitrate reductase from spinach (Spinacia

oleracea) leaves. Plant Physiol. 107, 451-457.

Magliery, T.J., Wilson, C.G.M., Pan, W., Mishler, D., Ghosh, I., Hamilton, A.D., and

Regan, L. (2005). Detecting protein-protein interactions with a green

fluorescent protein fragment reassembly trap: scope and mechanism. J. Am.

Chem. Soc. 127, 146-157.

Marchant, A., Kargul, J., May, S.T., Muller, P., Delbarre, A., Perrot-Rechenmann, C.,

and Bennett, M.J. (1999). AUX1 regulates root gravitropism in Arabidopsis by

facilitating auxin uptake within root apical tissues. EMBO J. 18, 2066-2073.

References 205

Marmagne, A., Ferro, M., Meinnel, T., Bruley, C., Kuhn, L., Garin, J., Barbier-Brygoo,

H., and Ephritikhine, G. (2007). A high content in lipid-modified peripheral

proteins and integral receptor kinases features in the Arabidopsis plasma

membrane proteome. Mol. Cell. Proteomics 6, 1980-1996.

Maudoux, O., Batoko, H., Oecking, C., Gevaert, K., Vandekerckhove, J., Boutry, M.,

and Morsomme, P. (2000). A plant plasma membrane H+-ATPase expressed in

yeast is activated by phosphorylation at its penultimate residue and binding of

14-3-3 regulatory proteins in the absence of fusicoccin. J. Biol. Chem. 275,

17762-17770.

May, T., and Soll, J. (2000). 14-3-3 proteins form a guidance complex with chloroplast

precursor proteins in plants. Plant Cell 12, 53-64.

McCabe, P.F., and Leaver, C.J. (2000). Programmed cell death in cell cultures. Plant

Mol. Biol. 44, 359-368.

Mhawech, P. (2005). 14-3-3 proteins - an update. Cell Res. 15, 228-236.

Minet, M., Dufour, M.-E., and Lacroute, F. (1992). Complementation of

Saccharomyces cerevisiae auxotrophic mutants by Arabidopsis thaliana cDNAs.

Plant J. 2, 417-422.

Moore, B.W., and Perez, V.J. (1967). Specific acidic proteins of the nervous system. In

Physiological and Biochemical Aspects of Nervous Integration, F. Carlton, ed

(Prentice Hall), pp. 343-359.

Moorhead, G., Douglas, P., Morrice, N., Scarabel, M., Aitken, A., and Mackintosh, C.

(1996). Phosphorylated nitrate reductase from spinach leaves is inhibited by

14-3-3 proteins and activated by fusicoccin. Curr. Biol. 6, 1104-1113.

Moorhead, G., Douglas, P., Cotelle, V., Harthill, J., Morrice, N., Meek, S., Deiting, U.,

Stitt, M., Scarabel, M., Aitken, A., and MacKintosh, C. (1999).

Phosphorylation-dependent interactions between enzymes of plant

metabolism and 14-3-3 proteins. Plant J. 18, 1-12.

Morsy, M., Gouthu, S., Orchard, S., Thorneycroft, D., Harper, J.F., Mittler, R., and

Cushman, J.C. (2008). Charting plant interactomes: possibilities and challenges.

Trends Plant Sci. 13, 183-191.

Mrowiec, T., and Schwappach, B. (2006). 14-3-3 proteins in membrane protein

transport. Biol. Chem. 387, 1227-1236.

Nair, R., and Rost, B. (2005). Mimicking Cellular Sorting Improves Prediction of

Subcellular Localization. J. Mol. Biol. 348, 85-100.

Nelson, B.K., Cai, X., and Nebenführ, A. (2007). A multicolored set of in vivo organelle

markers for co-localization studies in Arabidopsis and other plants. Plant J. 51,

1126-1136.

Nufer, O., and Hauri, H.-P. (2003). ER export: Call 14-3-3. Curr. Biol. 13, R391-R393.

References 206

Oecking, C., and Hagemann, K. (1999). Association of 14-3-3 proteins with the

C-terminal autoinhibitory domain of the plant plasma-membrane H+-ATPase

generates a fusicoccin-binding complex. Planta 207, 480-482.

Oecking, C., and Jaspert, N. (2009). Plant 14-3-3 proteins catch up with their

mammalian orthologs. Curr. Opin. Plant Biol. 12, 760-765.

Oecking, C., Eckerskorn, C., and Weiler, E.W. (1994). The fusicoccin receptor of plants

is a member of the 14-3-3 superfamily of eukaryotic regulatory proteins. FEBS

Lett. 352, 163-166.

Oecking, C., Piotrowski, M., Hagemeier, J., and Hagemann, K. (1997). Topology and

target interaction of the fusicoccin-binding 14-3-3 homologs of Commelina

communis. Plant J. 12, 441-453.

Ohad, N., Shichrur, K., and Yalovsky, S. (2007). The analysis of protein-protein

interactions in plants by bimolecular fluorescence complementation. Plant

Physiolology 145, 1090-1099.

Olsson, A., Svennelid, F., Ek, B., Sommarin, M., and Larsson, C. (1998). A

phosphothreonine residue at the C-terminal end of the plasma membrane

H+-ATPase is protected by fusicoccin-induced 14-3-3 binding. Plant Physiol. 118,

551-555.

Ottmann, C., Marco, S., Jaspert, N., Marcon, C., Schauer, N., Weyand, M.,

Vandermeeren, C., Duby, G., Boutry, M., Wittinghofer, A., Rigaud, J.-L., and

Oecking, C. (2007). Structure of a 14-3-3 coordinated hexamer of the plant

plasma membrane H+-ATPase by combining X-ray crystallography and electron

cryomicroscopy. Mol. Cell 25, 427-440.

Overmyer, K., Brosche, M., and Kangasjarvi, J. (2003). Reactive oxygen species and

hormonal control of cell death. Trends Plant Sci. 8, 335-342.

Palmgren, M.G. (2001). PLANT PLASMA MEMBRANE H+-ATPases: powerhouses for

nutrient uptake. Annu. Rev. Plant Physiol. Plant Mol. Biol. 52, 817-845.

Pan, S., Sehnke, P.C., Ferl, R.J., and Gurley, W.B. (1999). Specific interactions with TBP

and TFIIB in vitro suggest that 14-3-3 proteins may participate in the regulation

of transcription when part of a DNA binding complex. Plant Cell 11, 1591-1602.

Pandey, R., Muller, A., Napoli, C.A., Selinger, D.A., Pikaard, C.S., Richards, E.J.,

Bender, J., Mount, D.W., and Jorgensen, R.A. (2002). Analysis of histone

acetyltransferase and histone deacetylase families of Arabidopsis thaliana

suggests functional diversification of chromatin modification among

multicellular eukaryotes. Nucl. Acids Res. 30, 5036-5055.

Paul, A.-L., Sehnke, P.C., and Ferl, R.J. (2005). Isoform-specific subcellular localization

among 14-3-3 proteins in Arabidopsis seems to be driven by client interactions.

Mol. Biol. Cell 16, 1735-1743.

Paul, A.-L., Liu, L., McClung, S., Laughner, B., Chen, S., and Ferl, R.J. (2009).

Comparative interactomics: Analysis of Arabidopsis 14-3-3 complexes reveals

References 207

highly conserved 14-3-3 interactions between humans and plants. J. Proteome

Res. 8, 1913-1924.

Paul, C.B., and Russell, L.J. (2001). Cell death of barley aleurone protoplasts is

mediated by reactive oxygen species. Plant J. 25, 19-29.

Pendle, A.F., Clark, G.P., Boon, R., Lewandowska, D., Lam, Y.W., Andersen, J., Mann,

M., Lamond, A.I., Brown, J.W.S., and Shaw, P.J. (2005). Proteomic analysis of

the Arabidopsis nucleolus suggests novel nucleolar functions. Mol. Biol. Cell 16,

260-269.

Pennell, R.I., and Lamb, C. (1997). Programmed cell death in plants. Plant Cell 9,

1157-1168.

Piotrowski, M. (2008). Primary or secondary? Versatile nitrilases in plant metabolism.

Phytochemistry 69, 2655-2667.

Pracharoenwattana, I., Cornah, J.E., and Smith, S.M. (2005). Arabidopsis peroxisomal

citrate synthase is required for fatty acid respiration and seed germination.

Plant Cell 17, 2037-2048.

Preacher, K.J. (2001). Calculation for the chi-square test: an interactive calculation

tool for chi-square tests of goodness of fit and independence [Computer

software]. Available from http://www.quantpsy.org.

Purwestri, Y.A., Ogaki, Y., Tamaki, S., Tsuji, H., and Shimamoto, K. (2009). The 14-3-3

protein GF14c acts as a negative regulator of flowering in rice by interacting

with the florigen Hd3a. Plant Cell Physiol. 50, 429-438.

Repka, V., Fischerová, I., and Šilhárová, K. (2004). Methyl jasmonate is a potent

elicitor of multiple defense responses in grapevine leaves and cell-suspension

cultures. Biologia Plantarum 48, 273-283.

Reumann, S., Quan, S., Aung, K., Yang, P., Manandhar-Shrestha, K., Holbrook, D.,

Linka, N., Switzenberg, R., Wilkerson, C.G., Weber, A.P.M., Olsen, L.J., and Hu,

J. (2009). In-depth proteome analysis of Arabidopsis leaf peroxisomes

combined with in vivo subcellular targeting verification indicates novel

metabolic and regulatory functions of peroxisomes. Plant Physiol. 150,

125-143.

Riedel, J., Tischner, R., and Mack, G. (2001). The chloroplastic glutamine synthetase

(GS-2) of tobacco is phosphorylated and associated with 14-3-3 proteins inside

the chloroplast. Planta 213, 396-401.

Rienties, I.M., Vink, J., Borst, J.W., Russinova, E., and de Vries, S.C. (2005). The

Arabidopsis SERK1 protein interacts with the AAA-ATPase AtCDC48, the 14-3-3

protein GF14λ and the PP2C phosphatase KAPP. Planta 221, 394-405.

Rittinger, K., Budman, J., Xu, J., Volinia, S., Cantley, L.C., Smerdon, S.J., Gamblin, S.J.,

and Yaffe, M.B. (1999). Structural analysis of 14-3-3 phosphopeptide

complexes identifies a dual role for the nuclear export signal of 14-3-3 in ligand

binding. Mol. Cell 4, 153-166.

References 208

Roberts, M.R. (2003). 14-3-3 Proteins find new partners in plant cell signalling. Trends

Plant Sci. 8, 218-223.

Roberts, M.R., and Bowles, D.J. (1999). Fusicoccin, 14-3-3 proteins, and defense

responses in tomato plants. Plant Physiol. 119, 1243-1250.

Romeis, T., Ludwig, A.A., Martin, R., and Jones, J.D.G. (2001). Calcium-dependent

protein kinases play an essential role in a plant defence response. EMBO J. 20,

5556-5567.

Rosenquist, M. (2003). 14-3-3 proteins in apoptosis. Brazilian Journal of Medical &

Biological Research 36, 403-408.

Rosenquist, M., Alsterfjord, M., Larsson, C., and Sommarin, M. (2001). Data mining

the Arabidopsis genome reveals fifteen 14-3-3 genes. Expression is

demonstrated for two out of five novel genes. Plant Physiol. 127, 142-149.

Rosenquist, M., Sehnke, P., Ferl, R.J., Sommarin, M., and Larsson, C. (2000).

Evolution of the 14-3-3 protein family: Does the large number of isoforms in

multicellular organisms reflect functional specificity? J. Mol. Evol. 51, 446-458.

Saint-Jore-Dupas, C., Nebenfuhr, A., Boulaflous, A., Follet-Gueye, M.-L., Plasson, C.,

Hawes, C., Driouich, A., Faye, L., and Gomord, V. (2006). Plant N-glycan

processing enzymes employ different targeting mechanisms for their spatial

arrangement along the secretory pathway. Plant Cell 18, 3182-3200.

Sambrook, J., Fritsch, E.F., and Maniatis, T. (1989). Molecular Cloning: A Laboratory

Manual. (New York: Cold Spring Harbor Laboratory Press).

Schütze, K., Harter, K., and Chaban, C. (2009). Bimolecular Fluorescence

Complementation (BiFC) to study protein-protein interactions in living plant

cells. In Plant Signal Transduction, T. Pfannschmidt, ed (Humana Press), pp.

189-202.

Schoonheim, P.J., Veiga, H., da Costa Pereira, D., Friso, G., van Wijk, K.J., and de Boer,

A.H. (2007a). A comprehensive analysis of the 14-3-3 interactome in barley

leaves using a complementary proteomics and two-hybrid approach. Plant

Physiol. 143, 670-683.

Schoonheim, P.J., Sinnige, M.P., Casaretto, J.A., Veiga, H., Bunney, T.D., Quatrano,

R.S., and de Boer, A.H. (2007b). 14-3-3 adaptor proteins are intermediates in

ABA signal transduction during barley seed germination. Plant J. 49, 289-301.

Schultz, T.F., Medina, J., Hill, A., and Quatrano, R.S. (1998). 14-3-3 proteins are part

of an abscisic acid-VIVIPAROUS1 (VP1) response complex in the Em promoter

and interact with VP1 and EmBP1. Plant Cell 10, 837-847.

Sehnke, P.C., and Ferl, R.J. (2000). Plant 14-3-3s: omnipotent metabolic

phosphopartners? Sci. STKE 2000, pe1-.

Sehnke, P.C., Henry, R., Cline, K., and Ferl, R.J. (2000). Interaction of a plant 14-3-3

protein with the signal peptide of a thylakoid-targeted chloroplast precursor

References 209

protein and the presence of 14-3-3 isoforms in the chloroplast stroma. Plant

Physiol. 122, 235-241.

Sehnke, P.C., Chung, H.-J., Wu, K., and Ferl, R.J. (2001). Regulation of starch

accumulation by granule-associated plant 14-3-3 proteins. Proc. Natl. Acad. Sci.

USA 98, 765-770.

Sehnke, P.C., Laughner, B., Cardasis, H., Powell, D., and Ferl, R.J. (2006). Exposed loop

domains of complexed 14-3-3 proteins contribute to structural diversity and

functional specificity. Plant Physiol. 140, 647-660.

Shaner, N.C., Campbell, R.E., Steinbach, P.A., Giepmans, B.N.G., Palmer, A.E., and

Tsien, R.Y. (2004). Improved monomeric red, orange and yellow fluorescent

proteins derived from Discosoma sp. red fluorescent protein. Nat. Biotechnol.

22, 1567-1572.

Sheen, J. (2001). Signal transduction in maize and Arabidopsis mesophyll protoplasts.

Plant Physiol. 127, 1466-1475.

Shen, W., Clark, A.C., and Huber, S.C. (2003). The C-terminal tail of Arabidopsis

14-3-3ω functions as an autoinhibitor and may contain a tenth α-helix. Plant J.

34, 473-484.

Shikano, S., Coblitz, B., Wu, M., and Li, M. (2006). 14-3-3 proteins: regulation of

endoplasmic reticulum localization and surface expression of membrane

proteins. Trends Cell Biol. 16, 370-375.

Sinnige, M.P., Roobeek, I., Bunney, T.D., Visser, A.J.W.G., Mol, J.N.M., and de Boer,

A.H. (2005). Single amino acid variation in barley 14-3-3 proteins leads to

functional isoform specificity in the regulation of nitrate reductase. Plant J. 44,

1001-1009.

Sridha, S., and Wu, K. (2006). Identification of AtHD2C as a novel regulator of abscisic

acid responses in Arabidopsis. Plant J. 46, 124-133.

Staswick, P.E., Su, W., and Howell, S.H. (1992). Methyl jasmonate inhibition of root

growth and induction of a leaf protein are decreased in an Arabidopsis

thaliana mutant. Proc. Natl. Acad. Sci. USA 89, 6837-6840.

Su, W., Huber, S.C., and Crawford, N.M. (1996). Identification in vitro of a

post-translational regulatory site in the hinge 1 region of Arabidopsis nitrate

reductase. Plant Cell 8, 519-527.

Sugden, C., Donaghy, P.G., Halford, N.G., and Hardie, D.G. (1999). Two SNF1-related

protein kinases from spinach leaf phosphorylate and inactivate

3-hydroxy-3-methylglutaryl-coenzyme A reductase, nitrate reductase, and

sucrose phosphate synthase in vitro. Plant Physiol. 120, 257-274.

Sullivan, S., Thomson, C.E., Kaiserli, E., and Christie, J.M. (2009). Interaction

specificity of Arabidopsis 14-3-3 proteins with phototropin receptor kinases.

FEBS Lett. 583, 2187-2193.

References 210

Svennelid, F., Olsson, A., Piotrowski, M., Rosenquist, M., Ottman, C., Larsson, C.,

Oecking, C., and Sommarin, M. (1999). Phosphorylation of Thr-948 at the C

terminus of the plasma membrane H+-ATPase creates a binding site for the

regulatory 14-3-3 protein. Plant Cell 11, 2379-2392.

Taylor, N.J., and Fauquet, C.M. (2002). Microparticle bombardment as a tool in plant

science and agricultural biotechnology. DNA Cell Biol. 21, 963-977.

Toroser, D., Athwal, G.S., and Huber, S.C. (1998). Site-specific regulatory interaction

between spinach leaf sucrose-phosphate synthase and 14-3-3 proteins. FEBS

Lett. 435, 110-114.

Tzivion, G., Luo, Z., and Avruch, J. (1998). A dimeric 14-3-3 protein is an essential

cofactor for Raf kinase activity. Nature 394, 88-92.

Tzivion, G., Shen, Y.H., and Zhu, J. (2001). 14-3-3 proteins; bringing new definitions to

scaffolding. Oncogene 20, 6331-6338.

Ueki, S., Lacroix, B., Krichevsky, A., Lazarowitz, S.G., and Citovsky, V. (2008).

Functional transient genetic transformation of Arabidopsis leaves by biolistic

bombardment. Nat. Protocols 4, 71-77.

Ueno, Y., Ishikawa, T., Watanabe, K., Terakura, S., Iwakawa, H., Okada, K., Machida,

C., and Machida, Y. (2007). Histone deacetylases and ASYMMETRIC LEAVES2

are involved in the establishment of polarity in leaves of Arabidopsis. Plant Cell

19, 445-457.

Vain, P., Afolabi, A.S., Worland, B., and Snape, J.W. (2003). Transgene behaviour in

populations of rice plants transformed using a new dual binary vector system:

pGreen/pSoup. Theor. Appl. Genet. 107, 210-217.

van Hemert, M.J., Steensma, H.Y., and Heusden, G.P.H.v. (2001). 14-3-3 proteins: key

regulators of cell division, signalling and apoptosis. BioEssays 23, 936-946.

van Heusden, G.P.H., van der Zanden, A.L., Ferl, R.J., and Steensma, H.Y. (1996). Four

Arabidopsis thaliana 14-3-3 protein isoforms can complement the lethal yeast

bmh1 bmh2 double disruption. FEBS Lett. 391, 252-256.

Voinnet, O., Rivas, S., Mestre, P., and Baulcombe, D. (2003). An enhanced transient

expression system in plants based on suppression of gene silencing by the p19

protein of tomato bushy stunt virus. Plant J. 33, 949-956.

Würtele, M., Jelich-Ottmann, C., Wittinghofer, A., and Oecking, C. (2003). Structural

view of a fungal toxin acting on a 14-3-3 regulatory complex. EMBO J. 22,

987-994.

Waadt, R., Schmidt, L.K., Lohse, M., Hashimoto, K., Bock, R., and Kudla, J. (2008).

Multicolor bimolecular fluorescence complementation (mcBiFC) reveals

simultaneous formation of alternative CBL/CIPK complexes in planta. Plant J.

56, 505-516.

Walter, M., Chaban, C., Schütze, K., Batistic, O., Weckermann, K., Näke, C., Blazevic,

D., Grefen, C., Schumacher, K., Oecking, C., Harter, K., and Kudla, J. (2004).

References 211

Visualization of protein interactions in living plant cells using bimolecular

fluorescence complementation. Plant J. 40, 428-438.

Wang, A.H., Kruhlak, M.J., Wu, J., Bertos, N.R., Vezmar, M., Posner, B.I., Bazett-Jones,

D.P., and Yang, X.-J. (2000). Regulation of histone deacetylase 4 by binding of

14-3-3 proteins. Mol. Cell. Biol. 20, 6904-6912.

Wang, H., Qi, M., and Cutler, A.J. (1993). A simple method of preparing plant samples

for PCR. Nucl. Acids Res. 21, 4153-4154.

Watson, P., and Stephens, D.J. (2005). ER-to-Golgi transport: Form and formation of

vesicular and tubular carriers. Biochim. Biophys. Acta 1744, 304-315.

Weigel, D., and Glazebrook, J. (2002). Arabidopsis: a laboratory manual. (New York:

Cold Spring Harbor Laboratory Press).

Wilker, E.W., Grant, R.A., Artim, S.C., and Yaffe, M.B. (2005). A structural basis for

14-3-3σ functional specificity. J. Biol. Chem. 280, 18891-18898.

Witte, C.-P., Noël, L., Gielbert, J., Parker, J., and Romeis, T. (2004). Rapid one-step

protein purification from plant material using the eight-amino acid StrepII

epitope. Plant Mol. Biol. 55, 135-147.

Wittig, I., Braun, H.-P., and Schagger, H. (2006). Blue native PAGE. Nat. Protocols 1,

418-428.

Wroblewski, T., Tomczak, A., and Michelmore, R. (2005). Optimization of

Agrobacterium-mediated transient assays of gene expression in lettuce,

tomato and Arabidopsis. Plant Biotechnol. J. 3, 259-273.

Wu, K., Rooney, M.F., and Ferl, R.J. (1997a). The Arabidopsis 14-3-3 multigene family.

Plant Physiol. 114, 1421-1431.

Wu, K., Lu, G., Sehnke, P., and Ferl, R.J. (1997b). The heterologous interactions among

plant 14-3-3 proteins and identification of regions that are important for

dimerization. Arch. Biochem. Biophys. 339, 2-8.

Wu, K., Tian, L., Malik, K., Brown, D., and Miki, B. (2000). Functional analysis of HD2

histone deacetylase homologues in Arabidopsis thaliana. Plant J. 22, 19-27.

Xiao, B., Smerdon, S.J., Jones, D.H., Dodson, G.G., Soneji, Y., Aitken, A., and Gamblin,

S.J. (1995). Structure of a 14-3-3 protein and implications for coordination of

multiple signalling pathways. Nature 376, 188-191.

Xu, W.F., and Shi, W.M. (2006). Expression profiling of the 14-3-3 gene family in

response to salt stress and potassium and iron deficiencies in young tomato

(Solanum lycopersicum) roots: analysis by real-time RT-PCR. Ann Bot 98,

965-974.

Yaffe, M.B. (2002). How do 14-3-3 proteins work? Gatekeeper phosphorylation and

the molecular anvil hypothesis. FEBS Lett. 513, 53-57.

Yaffe, M.B., Rittinger, K., Volinia, S., Caron, P.R., Aitken, A., Leffers, H., Gamblin, S.J.,

Smerdon, S.J., and Cantley, L.C. (1997). The structural basis for

14-3-3:phosphopeptide binding specificity. Cell 91, 961-971.

References 212

Yang, Y., Costa, A., Leonhardt, N., Siegel, R., and Schroeder, J. (2008). Isolation of a

strong Arabidopsis guard cell promoter and its potential as a research tool.

Plant Methods 4, 6.

Yao, Y., Du, Y., Jiang, L., and Liu, J.-Y. (2007). Molecular analysis and expression

patterns of the 14-3-3 gene family from Oryza Sativa. J. Biochem. Mol. Biol. 40,

349-357.

Yoo, S.-D., Cho, Y.-H., and Sheen, J. (2007). Arabidopsis mesophyll protoplasts: a

versatile cell system for transient gene expression analysis. Nat. Protocols 2,

1565-1572.

Zhang, L., and Xing, D. (2008). Methyl jasmonate induces production of reactive

oxygen species and alterations in mitochondrial dynamics that precede

photosynthetic dysfunction and subsequent cell death. Plant Cell Physiol. 49,

1092-1111.

Zhong, S., Lin, Z., Fray, R., and Grierson, D. (2008). Improved plant transformation

vectors for fluorescent protein tagging. Transgenic Res. 17, 985-989.

Zhou, C., Labbe, H., Sridha, S., Wang, L., Tian, L., Latoszek-Green, M., Yang, Z., Brown,

D., Miki, B., and Wu, K. (2004). Expression and function of HD2-type histone

deacetylases in Arabidopsis development. Plant J. 38, 715-724.

Zimmermann, P., Hirsch-Hoffmann, M., Hennig, L., and Gruissem, W. (2004).

GENEVESTIGATOR. Arabidopsis microarray database and analysis toolbox.

Plant Physiol. 136, 2621 - 2632.

Zybailov, B., Rutschow, H., Friso, G., Rudella, A., Emanuelsson, O., Sun, Q., and van

Wijk, K.J. (2008). Sorting signals, N-terminal modifications and abundance of

the chloroplast proteome. PLoS ONE 3, e1994.

Appendices 213

Appendices

Appendix I: Media and Solutoins 214

Appendix I: Media and solutions

Solutions for plasmid minipreps (Sambrook et al., 1989)

Miniprep Solution I: 50 mM glucose, 10 mM EDTA, 25 mM Tris-HCl pH 8.0. Sterilise by

autoclaving.

Miniprep Solution II: 0.2 M NaOH, 1% SDS (freshly prepared).

Miniprep Solution III: 3 M sodium acetate, pH 4.7. Sterilise by autoclaving.

100% ethanol.

Sterile MQ H2O: autoclaved Milli-Q water.

Agarose gel electrophoresis

Agarose (Bioline).

1 x TAE running buffer (40 mM Tris-acetate, 1 mM EDTA). For convenience, make 50 x

TAE as a stock solution.

50x TAE (1L):

242 g Tris base

57.1 ml glacial acetic acid

100 ml 0.5 M EDTA (pH 8.0)

Orange G loading dye, 10x: 10 mM Tris-HCl, pH 8.0, 50 mM EDTA, 0.5% Orange G

(Sigma), 50% glycerol.

Ethidium bromide (1% solution, Applichem).

DNA ladder: Hyperladder I (Bioline).

Polymerase Chain Reaction (PCR)

BIOTAQ® DNA polymerase (Bioline).

10x NH4 Reaction Buffer (Bioline): 160 mM (NH4)2SO4, 670 mM Tris-HCl (pH8.8 at

25°C), 0.1% Tween-20.

MgCl2 stock solution (Bioline): 50 mM MgCl2.


dNTP mix (10 mM total): prepared from 100 mM stock solution of dATP, dCTP, dGTP

and dTTP (Bioline) by combining 2.5 mM of each.

Accuzyme® polymerase (Bioline).

10x AccuBuffer (Bioline): 600 mM Tris-HCl (pH8.3 at 25°C), 60 mM (NH4)2SO4, 100 mM

KCl, 20 mM MgSO4.

Chill-out 14® Liquid wax (MJ Research Inc.).

BigDye® DNA sequencing reaction

BigDye® Terminator v3.1 reaction mix [Applied Biosystems; purchased an aliquot from

Lotterywest Biomedical Facility: Genomics (LSBFG), Perth, Australia].

2.5x BigDye® sequencing buffer (Applied Biosystems, provided by LSBFG).

3 M sodium acetate, pH 4.6-4.7.

Preparation of competent E. coli cells

LB medium (1L): 10 g bacteriological tryptone, bacteriological yeast extract, 5 g NaCl,

pH 7.5.

LB plate: LB medium containing 15 g/L of bacteriological agar, autoclave sterilised.

LB medium containing 20 mM MgSO4.

TFB 1: 30 mM potassium acetate, 10 mM CaCl2, 50 mM MnCl2, 100 mM RbCl, 15%

glycerol. Adjust pH to 5.8 with 1 M acetic acid. 0.22 μm filter-sterilised.

TFB 2: 75 mM CaCl2, 10 mM RbCl, 15% glycerol, 10 mM MOPS or PIPES. Adjust pH to

6.5 with 1M KOH. 0.22 μm filter-sterilised.

Isolation of Arabidopsis thaliana mesophyll protoplasts

Protoplasting solution: 1% Cellulase (onozuka R-10; Duchefa Biochemie, Netherlands),

0.2% Macerozyme (Duchefa Biochemie), 0.4 M sucrose, 20 mM KCl, 10 mM CaCl2, 20

mM MES-KOH, pH 5.7; 0.45 μm filtered.

W5 solution: 154 mM NaCl, 125 mM CaCl2, 5 mM KCl, 2 mM MES-KOH, pH 5.7.


PS-1: A very gentle shaking motion (30 rpm) during the cell wall digestion process

helped in releasing protoplasts.

PS-2: a 75 μm nylon mesh was recommended to the use of sieving out undigested leaf

debris from the protoplasts by Yoo et al. (2007).

PEG-mediated transfection of A. thaliana mesophyll protoplasts

MaMg solution: 0.4 M mannitol, 15 mM MgCl2, 5 mM MES-KOH, pH 5.7

PEG-calcium solution: 40% (w/v) PEG 3350 (Sigma), 0.2 M mannitol, 100 mM CaCl2

PS: PEG4000 (Fluka, cat. no. 81240) was reported to perform an optimal protoplast

transfection efficiency (Yoo, et al. 2007)

Infiltration of Nicotiana benthamiana leaves

Acetosyringone (3, 5-Dimethoxy-4-hydroxy-acetophenone, Fluka, cat. No. 38766):

prepare 200 mM stock solution in DMSO.

Infiltration buffer: 10 mM MgCl2, 200 μM acetosyringone, 10 mM MES-KOH, pH 5.6.

Generation of transgenic Arabidopsis using floral dip transformation

5% sucrose (prepare freshly before use).

Silwet L-77 (Lehle Seeds, Cat# VIS-01).

Arabidopsis seed sterilisation

Anti-bacterial tablets (Milton): containing 50% sodium dichloroisocyanurate.

Seed sterilisation solution: resolve 0.0625 g of the ground up anti-bacterial tablet

powder in 1 ml deionised water, then add a drop of 1% Tween-20 and 9 ml of 95%

ethanol.

Screening Arabidopsis transformants

Basta® (200 g/L glufosinate ammonium, Hoechst, Australia).

Hygromycin B (100 mg/ml in H2O, A. G. Scientific Inc., USA).


1/2 MS medium plates: Murashige and Skoog (MS) basal salt mixture (Sigma, cat. No.

M5524) 2.15 g/L, 0.5% sucrose, 2 mM MES, pH 5.7, 0.8% agar.

PCR genotyping- a fast method

0.5 N NaOH

100 mM Tris, pH 8.0

Plant genomic DNA isolation -the CTAB method

Chloroform: Isoamyl alcohol (24:1, v/v)

CTAB extraction buffer:

1.4 M NaCl

100 mM Tris-HCl, pH 8.0

20 mM EDTA

2% (w/v) Ethyltrimethylammonium Bromide (CTAB)

1% (w/v) polycinylpyrollidine (PVP-40)

Heat the solution to dissolve and then autoclave.

Add β-Mercaptoethanol to 0.2% (v/v) before use.

RNase A solution (4 mg/ml, Invitrogen)

100% Isopropanol

Appendix II: Supplementary figures and tables 218

Appendix II: Supplementary figures and tables

Fig. S1. Strong background fluorescence adjacent to wound sites hindered evaluation of

BiFC experiments in infiltrated Arabidopsis leaves.

Epifluorescence and bright field images were captured from the same batch of agroinfiltrated

Arabidopsis leaves shown in Fig. 3-8. Strong background fluorescence was observed in the

cells surrounding the wounded leaf tissues. Scale bars = 50 μm.

Appendix II: Supplementary figures and tables 219

Table S1. Phenotypic variations of 14-3-3-split YFP carrying T1

transformants.

This table summarises aberrant phenotypes observed in eighteen 14-3-3-split YFP carrying T1

transformants. Observed phenotypes were grouped into ten categories for easier

comparisons between transformants.

Genotype T1 plant

Phenotypes

aberr

ant

leaf

morp

holo

gy

incre

ased

nu

mber

of ro

sett

e le

aves

serr

ate

d c

au

line leaves

aeria

l ro

sett

es o

n e

arly n

od

e

dela

y o

f flow

erin

g

(flo

ral bu

ds e

merg

ed >

10 w

after

sow

ing)

str

ong a

pic

al do

min

ance

(only

on

e p

rim

ary

ste

m)

inte

nsifie

d inflore

scence

bra

nches

short

ene

d s

tam

en f

ilam

ents

and

bro

wn

ish/im

ma

ture

anth

ers

short

ene

d s

iliq

ues o

r stu

nte

d s

iliqu

es

shru

nken s

ee

ds (

seed a

bort

ion)

epsilon∆-YN Ag29-1 x x x

epsilon∆-YC Ag20-1 x x x x

kappa-YN Ag31-7 x x x

Ag31-16 x x x x

kappa-YC

Ag32-1 x x

Ag32-9 x

Ag32-10 x

kappa∆-YC Ag34-1 x

Ag34-2 x x x x x

omega-YC Ag22-2 x x

omicron-YN Ag39-3 x x x x

Ag39-10 x x

omicron-YC Ag40-2 x

Ag40-7 x x

mu∆-YN Ag60-2 x x x

phi-YN Ag53-3 x x x

phi-YC Ag54-4 x x

Ag54-8 x

hung-chi liu - uwa research repository · a systematic in planta survey of 14-3-3 protein...

Documents