By T-DNA Mutant Analysis and Investigation of Molecular Interactions of Tandem Zinc Finger 1 (TZF1) in Arabidopsis Thaliana

Unraveling the Functions of Plant Ran GTPase-Activating Protein (RanGAP) by T-DNA Mutant Analysis and Investigation of Molecular Interactions of Tandem Zinc Finger 1 (TZF1) in Arabidopsis thaliana

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Thushani Rodrigo-Peiris, M.Eng. Graduate Program in Plant Cellular and Molecular Biology

The Ohio State University

2012

Dissertation committee:

Professor Iris Meier, Co-advisor

Professor Jyan-chyun Jang, Co-advisor

Professor Randy Scholl

Professor David Mackey

Professor Eric Stockinger

Thushani Rodrigo-Peiris

2012 ABSTRACT

RanGAP, the GTPase-Activating Protein (GAP) of the small GTPase Ran, is vital for nucleocytoplasmic trafficking of protein cargo in yeast and animals via the Ran cycle, and mitotic cell division. Arabidopsis thaliana contains two RanGAP paralogs, namely

RanGAP1 and RanGAP2. Despite the functional information of RanGAP revealed by extensive studies in yeast and animals, and its high degree of conservation in structure,

GAP activity and sub-cellular localization in plants, involvement of RanGAP in plant development is poorly understood. Studies in plants would provide clues on the evolutionary conservation and digression of RanGAP’s roles among eukaryotes, and would also enable insight into its effects on a whole-organism level in a system that undergoes indeterminate development. Here, I have used a genetic approach to generate a series of T-DNA insertion lines in Arabidopsis thaliana that contain decreasing RanGAP levels and increasing severity in vegetative and reproductive phenotypes. The mutants provide valuable experimental material to dissect the functions of plant RanGAP. I have analyzed several of these phenotypes in detail and revealed the importance of

Arabidopsis RanGAP in plant genesis, survival and development. The implications of mitotic involvement of RanGAP underlying these phenotypes points to a general conservation of RanGAP functionality in planta, in common with yeast and animals.

ii In an independent effort, molecular mechanisms underlying the functions of Tandem

Zinc Finger 1 of Arabidopsis thaliana (AtTZF1) were investigated. AtTZF1 is a plant-

unique CCCH-type tandem zinc finger (TZF) protein implicated in drought and cold

stress tolerance via regulation of gene expression. Studies with the most well studied human TZF; hTTP and AtTZF1 collectively suggest that the functions of AtTZF1 may be mediated by the dynamics in the assembly of stress responsive cytoplasmic mRNA- protein complexes; stress granules (SGs) and P-bodies (PBs), phosphorylation of AtTZF1 by the Mitogen-Activated Protein Kinase (MAPK) pathway and positive regulation of

ABA-mediated gene expression in planta. Understanding the mode of function of

AtTZF1 particularly in stress tolerance could be useful in developing agronomically important crops. To investigate the molecular mechanisms of AtTZF1 functions that currently remain elusive, assays on protein-protein interaction and AtTZF1 phosphorylation were conducted. Since the assays based on putative candidates were unable to reveal authentic molecular partners for AtTZF1, alternative approches may be used for future investigations.

iii

Dedicated to my parents, teachers, friends and family

iv ACKNOWLEDGEMENTS

I wish to express my utmost gratitude to my parents for being my first teachers, my inspiration, and for the unimaginable sacrifices they made on my behalf. Warmest thanks to my husband for being my friend, pillar of support and making me smile through the hardest times over the years. I am grateful to my brother and the rest of my family for being by my side always and unconditionally.

I am indebted beyond words to the kindness of the graduate school deans; Drs. Herness, Wallace and Slotnick. It was an honor and a priviledge to have received their guidance, encouragement and support in the time of need to complete my degree.

The work of my dissertation would not have been possible if not for the contributions of my advisors; Drs. Meier and Jang, and the dissertation committee; Drs. Scholl, Mackey and Stockinger. Special thanks are due to my collegues and friends in Rightmire Hall and Aronoff Laboratory for sharing their knowledge, ideas and materials, for insightful discussions and experimental help. Particularly I am grateful to my wonderful mentor Dr. Xianfeng Morgan Xu, and my collegues; Ms. Srimathi Bogamuwa, Dr. Nirodhini Siriwardana, Dr. Antje Feller, Dr. Siva Muthuswamy, Dr. Sowmya Venkatakrishnan, Dr. Srilakshmi Makkena, Ms. Jie Qu, Dr. Marcelo Pomeranz, Dr. Gireesha Mohannath, Ms. Veena Patil, Ms. Isabel Casas, Ms. Jamie Jackel, Dr. Ryuta Takeda, Dr. Qiao Zhao, Dr. Priya Raja, Ms. Katja Machemer, Dr. Sumire Fujiwara, Dr. Asuka Itaya and Dr. Kenneth Buckley. Your help, friendship and support over the years had inspired me beyond what you could possibly imagine.

v I am thankful to Ms. Horng-Jing (Heather) Wang and Mr. Li Zhang for their unpublished work cited in my dissertation. Assistance with microscopy by Mr. Brian Kemmenoe, Ms. Kathy Walken and Mr. Mike Shade, greenhouse support by Mr. Joe Takayama, technical support by Mr. Scott Hines and Mr. Dave Long, and numerous help by Ms. Melinda Parker, Ms. Laurel Shannon, Ms. Diane Furtney, Dr. Angela White, Dr. Sheila Westendorf, Ms. Karen Kyle and Ms. Lucinda Bolinger are greatly appreciated. Heartfelt thanks are also due to my supervisors at CLSE for constantly providing me a funding line to complete my degree and the exciting opportunities I received in teaching training.

I thank all those who were an inspiration, cheered me up, made me laugh, had fun with, got in trouble with, sent me wishes, gave a listening ear, fed me at hungry times, gave me smiles and hugs, and held my hand through all these years. Last but not least, thanks so much to OSU for this valuable and memorable experience. Go bucks!

vi VITA

2001 ……………………………………………B.Sc. in Botany University of Colombo, Sri Lanka

2004 ……………………………………………M.Eng. in Biotechnology Osaka University, Japan

2006 – present …………………………………Graduate Teaching/Research Associate Department of Molecular Genetics The Ohio State University

PUBLICATIONS

RanGAP1 is a continuous marker of the Arabidopsis cell division plane (2008). Xu, X.M., Zhao, Q., Rodrigo-Peiris, T., Brkljacic, J., He, C.S., Muller, S. and Meier, I. Proc Natl Acad Sci USA 105: 18637-18642

RanGAP is required for post-meiotic mitosis in female gametophyte development in Arabidopsis thaliana (2011). Rodrigo-Peiris, T., Xu, X.M., Zhao, Q., Wang, H-J. and Meier, I. J. Exp. Bot. 62(8): 2705-2714

FIELD OF STUDY

Plant Cellular and Molecular Biology

vii TABLE OF CONTENTS

Page

Abstract …………………………………………………………………………… ii

Dedication ………………………………………………………………………… iv

Acknowledgements……………………………………………………………….. v

Vita………………………………………………………………………………… vii

List of figures………………………………………………………………….…… xiv

List of tables……………………………………………………………………….. xvii

Chapters:

1. RanGAP is required for post-meiotic mitosis in Arabidopsis thaliana…………. 1

1.1 Abstract………………………………………………………… …………… 2

1.2 Introduction………………………………………………………………….. 3

1.3 Materials and Methods…………………………………………………….. .. 8

1.3.1 Plant material and growth conditions………………………………….. 8

1.3.2 PCR-based genotyping of T-DNA insertion lines……..………………. 8

1.3.3 Antibody preparation…………………………………………………… 9

1.3.4 Floral stage determination……………………………………………… 9

1.3.5 Ovule clearing and differential interference contrast (DIC) optics…….. 9

viii 1.3.6 Aniline blue staining…………………………………………………. .. 10

1.3.7 Cloning…………………………………………………………………. 10

1.3.8 Complementation assay……………………………………………….... 11

1.3.9 β-Glucuronidase assay…………………………………………………. 12

1.4 Results……………………………………………………………………….. 12

1.4.1 RanGAP single knock-out mutants lack observable phenotypes………. 12

1.4.2 RanGAP double knockout mutants rg1-1/rg1-1;rg2-3/rg2-3 were not obtained………………………………………………………………… 13

1.4.3 Transmission of the rg1-1 allele in the rg2-3 background is normal through the male parent but completely blocked through the female parent…… 14

1.4.4 rg1-1;rg2-3 ovules arrest during early megagametogenesis…………… 14

1.4.5 RanGAPs are expressed in the developing embryo sac and pollen……. 16

1.4.6 rg1-1/rg1-1;rg2-3/rg2-3 RanGAP knockout double mutants were rescued by complementation with RanGAP1 and RanGAP2 genomic constructs………………………………………………………………. 16

1.5 Discussion…………………………………………………………………… 17

1.5.1 RanGAP deficiency and lethality………………………………………. 17

1.5.2 RanGAP1 and RanGAP2 are functionally redundant in female gametophyte development……………………………………………………………. 21

1.5.3 Female gametophytes are arrested during a mitotic phase…………….. 21

1.5.4 Embryo sacs are affected more than the pollen………………………... 22

1.5.5 Outcomes of the study and future prospects…………………………… 24

2. Requirement of RanGAP for vegetative development and cell division in Arabidopsis thaliana……………………………………………………………… 44

2.1 Abstract………………………………………………………………………. 45

ix 2.2 Introduction…………………………………………………………………... 47

2.2.1 Ran cycle and the functions of RanGAP in animals and yeast………… 47

2.2.2 RanGAPs in Arabidopsis thaliana …………………………………….. 48

2.2.3 Shoot and root development in Arabidopsis thaliana …………………. 49

2.2.4 Cell division in plant vegetative meristems and the role of sugars…….. 51

2.2.5 Common phenotypes of cell cycle mutants and wild-type plants that were induced for cell division arrest…………………………………………. 54

2.3 Materials and Methods……………………………………………………….. 57

2.3.1 Plant material, growth conditions and constructs………………………. 57

2.3.2 PCR-based genotyping of T-DNA insertion lines……………………… 60

2.3.3 Immunoblot analysis…………………………………………………… 60

2.3.4 Embryo Observations…………………………………………………… 61

2.3.5 Scanning electron microscopy…………………………………………. 61

2.3.6 Sectioning……………………………………………………………… 62

2.3.7 Complementation assay………………………………………………… 62

2.3.8 β-Glucuronidase assay…………………………………………………. 63

2.3.9 Sucrose assay…………………………………………………………… 63

2.3.10 Confocal microscopy…………………………………………………… 64

2.4 Results……………………………………………………………………….. 64

2.4.1 RanGAP knockdown rg1-1/rg1-1;rg2-2/rg2-2 plants showed mild defects in vegetative growth and development…………………………………… 64

2.4.2 Severe RanGAP knockdown trans-heterozygous rg1-1/rg1-1;rg2-2/rg2-3 mutant seedlings showed prominent shoot and root phenotypes ……………………………………………………………… 65

x 2.4.3 rg1-1/rg1-1;rg2-2/rg2-3 showed adult vegetative and reproductive phenotypes...... 68

2.4.4 RanGAPs show high expression in the shoot and root meristems….….. 69

2.4.5 Shoot and root phenotypes were complemented with RanGAP1 and RanGAP2 genomic constructs…………………..…………………………………. 69

2.4.6 Phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 caused by depletion of RanGAP is likely independent from Ran cycle-based nuclear transport………….... 70

2.4.7 Cell division is restricted to compact regions in the shoot and root apices of rg1-1/rg1-1;rg2-2/rg2-3…………………………………………..…….. 71

2.4.8 Sucrose supplementation complements phenotypes of rg1-1/rg1-1;rg2-2/rg2- 3 seedlings ………………………...…………………………………… 72

2.4.9 Sucrose supplementation restores adult plant height and stature of rg1-1/rg1- 1;rg2-2/rg2-3…………………………………………………………… 73

2.4.10 Sucrose supplementation restores cell numbers and normal root morphology of rg1-1/rg1-1;rg2-2/rg2-3……………………………………………… 75

2.4.11 Sucrose complementation rescues cell division in the shoot and root apices of rg1-1/rg1-1;rg2-2/rg2-3 seedlings……………………………………… 75

2.5 Discussion……………………………………………………………………. 77

2.5.1 RanGAP, the cell cycle and vegetative plant development ……………. 77

2.5.2 RanGAP in shoot growth and development……………………………. 79

2.5.3 RanGAP in root growth and development……………………………… 81

2.5.4 Sucrose rescue of RanGAP mutant phenotypes ……………………….. 83

2.5.5 Future directions………………………………………………………… 86

3. Investigation of protein-protein interactions and stress response–related phosphorylation of tandem zinc finger 1 (TZF1) in Arabidopsis thaliana……… 127

3.1 Abstract………………………………………………………………………. 128

xi 3.2 Introduction………………………………………………………………….. 129

3.2.1 Zinc fingers, diversity and importance………………………………… 129

3.2.2 CCCH Zn finger proteins and CCCH-type Tandem Zn Finger proteins. 130

3.2.3 Cellular and Molecular mechanisms mediating TZF functions………… 130

3.2.4 Arabidopsis CCCH-type Tandem Zn Finger (TZF) proteins…………… 135

3.2.5 AtTZF1 structure, functional implications and stress response………… 136

3.3 Materials and Methods………………………………………………………... 139

3.3.1 Y-2-H screen……………………………………………………………. 139

3.3.2 Constructs………………………………………………………………. 140

3.3.3 Y-2-H assay…………………………………………………………….. 141

3.3.4 Transient protein expression in Nicotiana benthamiana ………………. 141

3.3.5 Microscopy…………………………………………………………...…. 142

3.3.6 Antibodies………………………………………………………..……… 142

3.3.7 Bioinformatics studies…………………………………………………... 143

3.3.8 Protein expression, purification and preparation for kinase assay……… 143

3.3.9 Kinase assay…………………………………………………………….. 145

3.4 Results…………………………………………………………………...……. 147

3.4.1 Evaluating putative AtTZF1 protein-protein interactions….…………… 147

3.4.1.1 Background………………………………………………………... 147

3.4.1.2 Y-2-H assay to test interaction of AtTZF1 with AtMPK11, AtRDL2 and AteIF2B……………………………………………………………. 149

3.4.2 Interactions of AtTZF1 with other putative candidates and self-interaction 149

3.4.2.1 Background……………………………………………………….. 149

xii 3.4.2.2 Y-2-H assay to test interactions of AtTZF1 with co-localized markers and self- interaction………………………..……………………… 151

3.4.3 Phosphorylation of AtTZF1 by AtMPK11……………………………… 151

3.4.3.1 Background……………………………………………………….. 151

3.4.3.2 AtTZF1 contains signature domains for MAPK-based phosphorylation, and dephosphorylation……………………………………………. 152

3.4.3.3 Protein expression and purification for AtTZF1 and AtMPK11….. 152

3.4.3.4 Optimization of the in vitro kinase- and phosphatase-assay protocols 154

3.4.3.5 Optimization of conditions for AtMPK11 kinase activity using MBP 155

3.4.3.6 Evaluation of phosphorylation of AtTZF1 by AtMPK11…………. 156

3.5 Discussion and future directions……………………………………………… 157

3.5.1 Interacting partners of AtTZF1…………………………………………. 157

3.5.2 Protein expression with respect to temperature………………………… 159

3.5.3 AtTZF1, AtMPK11 and phosphorylation……………………………… 161

Appendix A: GFP-AtAGO1 and GFP-AtMPK3 expression at 27°C and 5.5°C.. 180

References ………………………………………………………………………….. 181

xiii LIST OF FIGURES

Figure Page

1.1 Schematic representation of the gene structure and T-DNA insertions in RanGAP1 and RanGAP2………...……………………………………………………… 28

1.2 Immunoblot analysis confirms rg1-1 and rg2-3 as knockout (null mutant) lines 29

1.3 Siliques of heterozygous plants are semi-sterile……………………………... 30

1.4 RanGAP mutants are pre-fertilization lethal………………………………… 31

1.5 Floral, ovule and embryo sac development stages of Columbia correlate with those described for Landsberg erecta………………………………………………. 33

1.6 The majority of RanGAP mutant ovules are aborted at the two-nuclear embryo sac stage…………………………………………………………………………… 34

1.7 RanGAP promoter-GUS activity is detected in embryo sacs and pollen……… 35

1.8 Schematic representation of the genomic constructs used for complementation 36

1.9 Mutant lethality is rescued by genomic RanGAP constructs……………….… 37

2.1 T-DNA insert-position and protein expression of rg2-2 allele………………. 89

2.2 Seedling and adult plant phenotypes of the RanGAP knockdown rg1-1/rg1-1;rg2- 2/rg2-2 mutant………………………………………………………………… 90

2.3 Reproductive phenotypes of the RanGAP knockdown rg1-1/rg1-1;rg2-2/rg2-2 mutant and phenotypic complementation……………………...……………… 92

2.4 Embryo observations and molecular characterization of rg1-1/rg1-1;rg2-2/rg2- 3 mutant………………………………………………………………………... 94

2.5 Seedling phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 mutant…………….…..……. 96

xiv 2.6 Scanning electron micrographs show delayed shoot apex development in rg1-1/rg1- 1;rg2-2/rg2-3 seedlings...... 98

2.7 Development of the first pair of true leaves is abnormal in rg1-1/rg1-1;rg2-2/rg2-3 seedlings and successive initiation of true leaves in a radial pattern is perturbed ……………………………………………………………………... 99

2.8 Histological analysis reveals disorganized cell arrangement in the developing true leaves and shoot apex of rg1-1/rg1-1;rg2-2/rg2-3…………………………… 100

2.9 Severely affected rg1-1/rg1-1;rg2-2/rg2-3 seedlings are lethal. Adventitioius shoot meristems of surviving rg1-1/rg1-1;rg2-2/rg2-3 seedlings give rise to a compact shoot with multiple rosettes…………………………………………………… 102

2.10 Root morphological and anatomical phenotypes of rg1-1/rg1-1;rg2-2/rg2-3.. 104

2.11 Adult plant phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 mutant……………..…… 106

2.12 Microarray expression data shows enhanced RanGAP expression in the shoot apex and root meristem……………………………………………………………. 107

2.13 Assay for RanGAP promoter-GUS expression shows a high level of staining in the vegetative meristems of shoots and roots……………………………………. 108

2.14 Phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 are rescued by native promoter-driven genomic constructs…………………………………..………………………. 109

2.15 Nuclear import is not impeded in rg1-1/rg1-1;rg2-2/rg2-3…………………. 111

2.16 CycB1;1::GUS expression is restricted to smaller regions in the shoot and root apices of rg1-1/rg1-1;rg2-2/rg2-3 compared to rg1-1/rg1-1;RG2/rg2-2…... 113

2.17 Growth and developmental phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 seedlings are rescued by sucrose supplementation. ………………………………...……… 114

2.18 Adult shoot phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 are rescued by sucrose supplementation………………………………………...…………………….. 116

2.19 Cell numbers at the root tip and root morphology are rescued by sucrose supplementation..….………………………………………………………….. 119

2.20 Sucrose supplementation rescues CycB1;1::GUS expression in the shoot apex of rg1-1/rg1-1;rg2-2/rg2-3……………...………………………………………. 122

xv 2.21 Sucrose supplementation rescues CycB1;1::GUS expression in the root apex of rg1- 1/rg1-1;rg2-2/rg2-3…………………………………………………………… 124

3.1 Y-2-H assay failed to show a direct interaction for AtTZF1-AtMPK11, AtTZF1- AtRDL2 and AtTZF1-AteIF2B using full-length clones...... 163

3.2 Y-2-H assay failed to show interactions between AtTZF1 and PB/SG markers, and AtTZF1 self-interaction…………………………………………………..…… 166

3.3 Sequence analysis revealed MAPK-docking, phosphorylation and phosphatase- binding sites in the AtTZF1 protein…………………………………………... 168

3.4 Cold room conditions increased protein levels of N. benthamiana leaves expressing GFP-AtTZF1…………………………………………………………………. 169

3.5 Purification of GFP-AtMPK11 and AtTZF1-GST for in vitro kinase assay….. 170

3.6 GFP-AtMPK11 and free GFP were prepared concurrently for comparative in vitro kinase assays………………………………………………………………..…. 172

3.7 in vitro kinase assay detected phosphorylation/de-phosphorylation events under the experimented conditions..……………………………………………….… 173

3.8 Magnetic bead-bound GFP-AtMPK11 is able to phosphorylate MBP………… 175

3.9 MBP phosphorylation by AtMPK11 was improved via modifications of the in vitro kinase assay protocol…………………………………………………………. 177

3.10 AtMPK11 phosphorylates MBP, but not AtTZF1, in the in vitro kinase assay.. 179

Appendix A: GFP-AtAGO1 and GFP-AtMPK3 expression at 27°C and 5.5°C… 180

xvi LIST OF TABLES

Table Page

1.1 Primers used in this study………………………………………………………….38

1.2 Floral stages, landmark characteristics and female gametophyte development…...39

1.3 Expected and observed F3 segregation ratios derived from a cross between rg1- 1/rg1-1;RG2/RG2 and RG1/RG1;rg2-3/rg2-3…………………………………..…40

1.4 Number of aborted ovules from rg1-1/rg1-1;RG2/rg2-3 and wild-type (Col) siliques……………………………………………………………………………..41

1.5 Transmission of the rg1-1;rg2-3 genotype through the male and female gametophytes…………………………………………………...………………….42

1.6 Percentage of female gametophytes at selected stages of development in the siliques of semi-sterile heterozygote, rg1-1/rg1-1;RG2/rg2-3…………………………..…43

2.1 Silique data from rg1-1/rg1-1;RG2/rg2-2 and rg1-1/rg1-1;rg2-2/rg2-3 plants.…125

2.2 Transmission of the rg1-1;rg2-3 genotype in comparison to rg1-1;rg2-2 through the male gametophyte…………………………………………………….…...….126

xvii CHAPTER 1

RANGAP IS REQUIRED FOR POST-MEIOTIC MITOSIS IN FEMALE GAMETOPHYTE DEVELOPMENT IN ARABIDOPSIS THALIANA

1 1.1 Abstract

RanGAP is the GTPase-activating protein of the small GTPase Ran and is involved in nucleocytoplasmic transport in yeast and animals via the Ran cycle and in mitotic cell division. Arabidopsis thaliana has two copies of RanGAP which are ~60% identical at the protein level, RanGAP1 and RanGAP2. To investigate the function of plant RanGAPs on a whole-organism level, T-DNA insertional mutants were analyzed. Arabidopsis plants with a knockout allele for either RanGAP1 or RanGAP2 lacked an observable phenotype. Analysis of the segregating progeny showed that double mutants in RanGAP1 and RanGAP2 are female gametophyte defective. Ovule clearing with differential interference contrast (DIC) optics showed that mutant female gametophytes were arrested at the interphase, predominantly after the first mitotic division following meiosis. In contrast, mutant pollen developed and functioned normally as seen by analysis of genetic crosses, observation of pollen morphology by light microscopy and pollen tube transmission in aniline blue-stained pistils. These results show that the RanGAPs are redundant and indispensable for female gametophyte development in Arabidopsis. Nuclear division arrest during a mitotic stage suggests a role for plant RanGAP in cell cycle progression in mitotic cell division during female gametophyte development.

2 1.2 Introduction

RanGAP (RG) is the GTPase-Activating Protein (GAP) of the small GTPase Ran and is conserved in eukaryotes including yeast, animals and plants (Hopper et al., 1990; Melchior et al., 1993; DeGregori et al., 1994; Bischoff et al., 1995; Meier, 2000; Pay et al., 2002). It is involved in nucleocytoplasmic transport of proteins that contain nuclear localization signals and/or nuclear export signals via the Ran cycle in yeast and animals (Bischoff et al., 1995; Traglia et al., 1996; Hutten et al., 2008) and in mitotic cell division (Bamba et al., 2002; Kusano et al., 2004). Roles of the Ran cycle in animal mitosis include cell cycle progression, spindle assembly, centrosome duplication, chromosome alignment, segregation and decondensation, as well as nuclear envelope re-assembly following mitosis (Ren et al., 1994; Hetzer et al., 2000; Zhang and Clarke, 2000; Gruss et al., 2001; Zhang et al., 2002; Arnaoutov and Dasso, 2003; Arnaoutov and Dasso, 2005; Ciciarello et al., 2007). In yeast and animals, null mutants of RanGAP homolog are lethal (Hartwell, 1967; DeGregori et al., 1994; Bamba et al., 2002; Kusano et al., 2004).

Arabidopsis thaliana carries two paralogs of RanGAP, RanGAP1 (RG1) and RanGAP2 (RG2) which show ~60% identity with each other and ~20% identity with either the Saccharomyces cereviceae homolog (ScRna1p) or the human RanGAP homolog (HsRanGAP) (Rose and Meier, 2001). They complement the yeast temperature sensitive rna1p mutant, rna1-1 (Pay et al., 2002), thus confirming conserved GAP (i.e. GTPase- Activating Protein) activity in plant RanGAPs. Publicly available microarray data sets at GENEVESTIGATOR (www.genevestigator.ethz.ch; Zimmermann et al., 2004) show a high level of expression for both RanGAPs in regions with high mitotic activity, including shoot apex and root meristem, in addition to basal level of expression in almost all tissues. RanGAP1 expression is also reported as high in both ovules and pollen, while RanGAP2 expression is high in ovules but very low in pollen.

3 As in animal cells, Arabidopsis RanGAP1 and RanGAP2 are located at the nuclear envelope in differentiated cells and during interphase of cycling cells in root tip (Rose and Meier, 2001; Pay et al., 2002; Jeong et al., 2005). Several of the mitotic locations of RanGAP in animals are also conserved in Arabidopsis, such as a concentration at the kinetochores and at the spindle midzone (Matunis et al., 1998; Joseph et al., 2002). These localization data suggest a possibility of conserved functions of RanGAP in nuclear transport and cell division in plants as well. In addition, Arabidopsis RanGAP1 is a continuous marker of the cell division plane, found concentrated at the preprophase band and retained at the cortical division zone, the phragmoplast and the growing edge of the cell plate (Pay et al., 2002; Jeong et al., 2005; Xu et al., 2008). However, studies investigating the functional significance of RanGAP in plants have been limited. Inducible knockdown experiments indicate a role for Arabidopsis RanGAPs in proper cell plate establishment and positioning during cytokinesis (Xu et al., 2008). In addition, RanGAP2 was found to be involved in disease resistance to potato virus X (PVX) via an unknown mechanism (Sacco et al., 2007; Tameling and Baulcombe, 2007).

The life cycle of an angiosperm alternates between a long-lived diploid sporophyte generation and a short-lived haploid male/female gametophyte generation (Drews et al., 1998; Drews and Yadegari, 2002). Both gametophytes, pollen (male) and embryo sac (female), result from a conserved program of cell division starting from a diploid sporogenous initial cell namely microspore- (male) and megaspore-(female) mother cell. Following a meiotic division of the mother cell, each resulting microspore undergoes two mitotic divisions to form a male gametophyte (pollen grain) consisting of a vegetative cell and two sperm cells. Out of the four daughter cells produced by meiosis of the megaspore mother cell, the three distal cells undergo programmed cell death and the persisting proximal cell, the megaspore, undergoes three mitotic divisions to form a female gametophyte (embryo sac) containing eight haploid nuclei. The two nuclei resulting from the first mitosis of the megaspore separates by vacuole formation, after which the two subsequent mitotic divisions of each nucleus takes place at the two poles

4 resulting in a syncytium. One nucleus from each pole (called polar nuclei) migrates to a central location, which following cytokinesis/cellularization of the embryo sac, fuse to form the diploid central cell nucleus (called secondary nucleus). Near the distal pole (micropylar end), the three cells each carrying a haploid nucleus become differentiated to form the two synergids and an egg cell which together with the three antipodal cells at the proximal end (chalazal end) constitute the seven-nuclear seven-celled embryo sac which is surrounded by several layers of sporophytic tissue; the integuments. The embryo sac together with the sporophytic tissue that surrounds it constitutes the ovule. Fertilization of the haploid male gamete (a sperm cell) and the female gamete (the egg cell) gives rise to the diploid zygote to begin the sporophytic generation of the life cycle (Reviewed in Drews et al., 1998; Drews and Yadegari, 2002; Yadegari and Drews, 2004).

Mutant studies are greatly facilitated by the haploid nature of the gametophytes because they could help study the functions of essential genes (i.e. genes that are indispensable to support cellular life), whose mutations are difficult to investigate in the sporophyte (Drews and Yadegari, 2002; Shi et al., 2005). Thus, study of the haploid phase appears a promising approach to dissect the functions of RanGAP in plants, an essential gene which is known to be indispensable for fundamental cellular processes [i.e. mitosis and nucleocytoplamic transport in yeast and animals - Bischoff et al., 1995; Traglia et al., 1996; Bamba et al., 2002; Kusano et al., 2004; Meier, 2005; Hutten et al., 2008], which in homozygous mutant condition may be lethal in plants as well. Although both forward and reverse genetic approaches have been utilized for identification of essential gene functions in the gametophytes, forward genetics have been hampered by functional redundancy, thus leaving reverse genetics an attractive option for these proteins. Arabidopsis RanGAP1 and RanGAP2 that show a high degree of similarity between them could be expected to show redundancy in at least some of their functions which may hinder the application of forward genetics to study their functions. In gametophytic mutant studies, although commonly both gametophytes are affected to some degree by the disruption of essential genes given their likely common roles in both systems, studies

5 have shown that what looks like a general pathway could affect one gametophyte more than the other or simply one gametophyte alone (Drews et al., 1998; Siddiqi et al., 2000; Huanca-Mamani et al., 2005; Shi et al., 2005).

On the other hand, understanding of gametophyte development and functioning is important given the importance of the haploid life cycle in plant development. Little is known about the genetic control and molecular mechanisms of gametogenesis in higher plants. Although many mutations defective in female and male gametophytic functions have been isolated by genetic screens (Feldmann et al., 1997; Christensen et al., 1998; Drews et al., 1998; Drews et al., 1998; Yang and Sundaresan, 2000; Johnson et al., 2004; Shi et al., 2005), only a few of these genes have been characterized (Shi et al., 2005). Due to the unique development, structure and function of the gametophytes compared to that of the sporophyte and the differences between male and female gametophytes themselves, it remains particularly important to identify their molecular players in order to evaluate how they are networked to bring about their uniqueness. Thus identifying RanGAPs’ roles in the male and female gametophytes would add to our understanding of how these unique systems work.

A high proportion of the characterized female gametophytic mutants are impaired during the mitotic cell cycle progression (Brukhin et al., 2005; Shi et al., 2005) suggesting that mitosis is a critical stage in female gametophyte development. Among these, a significant representation of genes encoding essential cell division-related proteins is evident. Mutants of positive regulators of cell cycle have resulted in the arrest of the embryo sac nuclei at various mitotic stages. These mutants include defects in PROLIFERA (homolog of yeast Mcm7 important for DNA replication in S phase) (Springer et al., 1995; Springer et al., 2000), INCURVATA2 (putative catalytic subunit of DNA polymerase α) (Barrero et al., 2007), mutants of subunits of the anaphase-promoting complex/cyclosome (APC/C) including NOMEGA (Capron et al., 2003; Kwee and Sundaresan, 2003), STIMPY (a homeobox protein important for cell cycle progression at G2 phase) (Wu et al., 2005; Wu

6 et al., 2007), γ-Tubulin (important for assembly of spindle, phragmoplast and cortical microtubule arrays) (Binarova et al., 2006; Pastuglia et al., 2006), SLOW WALKER1 (a protein involved in 18S Ribosomal RNA biogenesis) (Shi et al., 2005) and chromatin- remodeling protein 11 (CHR11) (Huanca-Mamani et al., 2005), which are all likely involved in cell division-specific functions. When plants are deficient in EXPORTIN1 (XPO1/CRM1); a protein that is involved in the nucleocytoplasmic transport and mitotic cell division via the Ran cycle, the mutants show less pronounced male gametophyte defects (Arnaoutov et al., 2005; Blanvillain et al., 2008). On the other hand, mutants of negative cell cycle regulators have commonly developed supernumerary nuclei [Lin, 1978; Ebel et al., 2004 and mutants of FOUR LIPS (FLP); Makkena, S. and Lamb, R. Department of Molecular Genetics. Personal communication] or more than seven cells in the embryo sac [mutants of FOUR LIPS (FLP); Makkena, S. and Lamb, R. Department of Molecular Genetics. Personal communication]. Essential genes involved in general cellular processes unrelated to cell division have also shown arrests in mitotic stages of female gametophyte development. In this regard, female gametophyte development has shown to be sensitive to the availability of a large number of genes that are crucial for general cellular functions such as transcriptional regulation, signal transduction, protein degradation and secondary metabolism (Pagnussat et al., 2005). However, particularly owing to the inaccessible nature of the female gametophytes which are embedded within the sporophytic tissues of the mother plant, further molecular and biochemical dissection in the female gametophytes to identify the exact underlying cause for arrest had been particularly challenging (Howden et al., 1998; Drews and Yadegari, 2002).

In an approach to identify functional roles of RanGAP in planta on a whole-organism level and to gain insight to its mode of function, T-DNA mutants of RanGAP were generated and studied. Here, genetic and cell-biological evidence was generated using a RanGAP knockout mutant to demonstrate that RanGAP is essential for female gametophyte development in Arabidopsis and that RanGAP1 and RanGAP2 are redundant for this function. Female gametophytes containing RanGAP double null

7 mutant alleles were arrested post-meiosis, predominantly at the two-nuclear mitotic stage thereby suggesting a role for RanGAP in cell cycle progression in plant mitosis. In contrast to female gametophyte arrest, the mutant pollen produced by the same mother plants developed and functioned normally.

1.3 Materials and Methods

1.3.1 Plant material and growth conditions

This study was based on RanGAP1 (At3g63130) and RanGAP2 (At5g19320) of Arabidopsis thaliana. T-DNA insertion mutant rg1-1 (SALK_058630) in Columbia (Col) ecotype was acquired from the Arabidopsis Biological Resource Center (ABRC). T-DNA mutant rg2-3 (FLAG_184A06) in Wassilewskija (Ws) was acquired from the Versailles T-DNA lines collection (Bechtold et al., 1993; Bouchez et al., 1993). The position of the T-DNA insert in rg1-1 and rg2-3 alleles were confirmed by sequencing to lie ~720 bp downstream and ~190 bp upstream of the start codon of RanGAP1 and RanGAP2 respectively. An F1 generation was obtained from a cross between the two mutants and was allowed to self-fertilize to obtain the F2 generation. Work leading to the production of F2 generation was conducted by Xu, X.M. (Xu, 2007). Arabidopsis seedlings were

grown in soil under standard long-day conditions (16 h day/ 8 h night).

1.3.2 PCR-based genotyping of T-DNA insertion lines

For PCR-based genotyping of T-DNA insertion lines, genomic DNA was extracted as described (Krysan et al., 1999). All primer sequences are summarized in Table 1.1. For rg1-1 and rg2-3 screening, primer combinations RanGAP5.2/ LBa1 and 032721FP/ LBwas were used respectively.

8 1.3.3 Antibody preparation

Anti-RanGAP1 antibody has been described in (Jeong et al., 2005). For the anti- RanGAP2 antibody, full length RanGAP2 protein was expressed as a His-tag fusion protein using the pDEST17 vector (Invitrogen). After purification of the recombinant protein with a Ni-NTA resin column and excision from a preparative SDS/PAGE gel, a guinea pig antiserum (OSU-GP4) was produced by Cocalico Biologicals (Reamstown, PA).

1.3.4 Floral stage determination

Definitions for floral and ovule development stages used by Smyth et al., 1990 and Schneitz et al., 1995 for Landsberg erecta (Ler) ecotype were adopted in this study (Table 1.2) because wild-type Columbia (WT Col) and wild-type Wassilewskija (WT Ws) also showed a similar developmental pattern (Figure 1.5 and data not shown). Denotations 12- early, 12-mid and 12-late stages refer to when a) petals align with anthers of long stamens, b) petals are midway between anthers of long stamens and stigma, and c) petals are at the level with the stigma respectively. Stage 13-early refers to when buds open with petals clearly visible above the sepals but anthesis is yet to occur and stamens lie below the level of the stigma. Stage 14-early was determined by pre-experimentation at ‘13 h after anthers of flowers at the stage of 13-early have been emasculated’, ensuring that mature embryo sacs could be halted at the eight-nuclear stage without fertilization. ‘Early’ refers to the beginning of the floral stage showing hallmark features of the corresponding stage (Table 1.2).

1.3.5 Ovule clearing and differential interference contrast (DIC) optics

Dissected pistils were fixed in absolute ethanol : acetic acid 9:1 (v/v) at 4°C overnight and washed with 90% ethanol for 1 h at room temperature. Pistils were subsequently

9 washed with 70% ethanol for 1 h at room temperature and cleared with a clearing solution (chloral hydrate 8 g: Glycerol 1 ml: water 2 ml) overnight at room temperature.

1.3.6 Aniline blue staining

Pistils were fixed for 2 h at room temperature in glacial acetic acid : absolute ethanol 1:3 (v/v) and submerged in 1 M NaOH overnight at 4°C for softening of the tissue. Tissues were washed with 50 mM potassium phosphate buffer (pH 7.5) and the ovules were exposed by dissection of the carpel wall. Pistils were stained with 0.1% aniline blue for 4-6 h and viewed under UV light.

1.3.7 Cloning

For the RG1 promoter-GUS construct, a ~1.2 kb genomic fragment upstream of the coding region of RanGAP1 was PCR-amplified using Taq DNA Polymerase (New England Biolabs) from a genomic DNA preparation (DNeasy plant mini kit, Qiagen) using primers MLU1 and Promoter 3’ in Table 1.1. The insert was confirmed by sequencing and contained a ~0.73 kb of the annotated promoter, 5’UTR region and the intron of RanGAP1 (illustrated in Figure 1.8A) which was subsequently sub-cloned into PCR2.1-TOPO TA-cloning vector (Invitrogen). A BamHI- and Not1-restriction enzyme digested fragment from the recombinant pCR2.1-TOPO vector containing the RanGAP1 promoter was cloned into pENTR 3C (Invitrogen) and moved to pMDC162 (Curtis and Grossniklaus, 2003) by LR recombination reaction (Invitrogen). RG2 promoter-GUS construct containing a ~0.85 kb genomic fragment upstream of the coding region of RanGAP2 was cloned the same way using primers Pro.RanGAP2 F and Pro.RanGAP2 R in Table 1.1. For sub-cloning from PCR2.1-TOPO TA-cloning vector into pENTRTM 3C, the same restriction enzymes BamHI and NotI were used. RG1 promoter-GUS and RG2 promoter-GUS constructs were generated by Wang, H-J.

10 The native promoter-RG1 construct (Figure 1.8A) was generated by cloning a ~2.2 kb fragment of RanGAP1 into the RG1 promoter-GUS construct described above. The ~2.2 kb genomic fragment consisting of the coding region of RanGAP1, the 3’UTR and a ~300 bp intergenic region downstream of the 3’UTR was amplified from a genomic DNA preparation (DNeasy plant mini kit, Qiagen) using Platinum Pfx DNA polymerase (Invitrogen) and primers RG1-F1-CACC (AscI) and RG1-intergenic R1 (AscI) in Table 1.1. The insert was sub-cloned into pENTR/D-TOPO (Invitrogen). After sequence confirmation, the AscI-cut ~2.2 kb genomic fragment was recombined into the AscI-cut RG1 promoter-GUS construct by traditional ligation.

The native promoter-RG2 construct (Figure 1.8B) was generated by cloning a ~4 kb fragment of RanGAP2 genomic DNA into pGWB1 binary vector (The pGWB1 binary vector was a kind gift of Dr.Tsuyoshi Nakagawa-Research Institute of Molecular Genetics, Shimane University). The ~4 kb fragment containing ~1.2 kb of annotated native RanGAP2 promoter, 5’UTR region, the intron, the coding region and a ~200 bp 3’UTR was amplified from a genomic DNA preparation (DNeasy plant mini kit, Qiagen) using Platinum Pfx DNA polymerase (Invitrogen) and primers RG2-PDONR-4KB-F and RG2-PDONR-4KB-F in Table 1.1. and sub-cloned into pDONR 221 (Invitrogen) by BP recombination cloning (Invitrogen). The insert was confirmed by sequencing and subsequently introduced into pGWB1 by LR recombination reaction (Invitrogen).

1.3.8 Complementation assay

Plasmids carrying native promoter-RG1/pMDC162 and native promoter-RG2/pGWB1 were mobilized into the Agrobacterium strain GV3101 by electroporation. Subsequently, Arabidopsis rg1-1/rg1-1;RG2/rg2-3 plants were transformed by floral dipping (Clough and Bent, 1998) and the primary transformants (T1 generation) were selected by Hygromycin resistance (30 µg/ml). T1 and T2 generations were screened by PCR genotyping for rescued rg1-1/rg1-1;rg2-3/rg2-3 plants.

11 1.3.9 β-Glucuronidase assay

Histochemical staining for GUS activity was performed as described in Jefferson et al., 1987. In short, tissues were fixed for 1 h in 90% acetone at -20°C, rinsed with 0.2 M

Na2HPO4, stained with 1 mM 5-bromo-4-chloro-3-indolyl-β-D-glucuronic acid (X-Gluc) in 50 mM potassium phosphate buffer (pH 7.0) and cleared with 70% and 90% ethanol (v/v). Degree of staining of ovules was modulated by manipulating the duration of staining. Ovules were cleared with a chloral hydrate solution (chloral hydrate 8 g: Glycerol 1 ml: water 2 ml) and observed with DIC optics.

1.4 Results

1.4.1 RanGAP single knock-out mutants lack observable phenotypes

To investigate the roles of RanGAP in Arabidopsis by the study of mutant phenotypes, single RanGAP1 and RanGAP2 T-DNA null mutants were analyzed. No observable phenotypes were detected in T-DNA single mutants for RanGAP1; SALK_058630 (rg1-1) and RanGAP2; FLAG_184A06 (Wassilkja) (rg2-3). Phenotypic analysis of the single mutants was carried out by Xu, X.M. and Zhao, Q. Figure 1.1 shows the gene structure of RanGAPs and T-DNA insertion maps of rg1-1 and rg2-3 alleles. rg1-1 and rg2-3 were confirmed as knockout alleles by Western blot analysis (Figure 1.2). The single mutants rg1-1/rg1-1;RG2/RG2 and RG1/RG1;rg2-3/rg2-3 were crossed to obtain the F1 progeny for segregation analysis in F2 and F3 generations.

12 1.4.2 RanGAP double knockout mutants rg1-1/rg1-1;rg2-3/rg2-3 were not obtained

The segregation of the F2 progeny of the cross between single RanGAP mutants; rg1- 1/rg1-1;RG2/RG2 and RG1/RG1;rg2-3/rg2-3 yielded no double knockout mutants indicating lethality and redundancy of RanGAP1 and RanGAP2 for the underlying function. RG1/rg1-1;rg2-3/rg2-3 and rg1-1/rg1-1;RG2/rg2-3 plants were identified by PCR-based genotyping in the F2 generation. They had no observable vegetative phenotype, suggesting that a reduction of the overall RanGAP amount to ~25% is sufficient for growth and development under standard laboratory conditions. RG1/rg1- 1;rg2-3/rg2-3 and rg1-1/rg1-1;RG2/rg2-3 F2 plants were allowed to self-fertilize and the F3 segregation ratio was determined by PCR-based genotyping. Absence of double knockout mutants was confirmed in the F3 generation, accompanied by non-Mendelian ratios of segregation (Table 1.3). The segregation ratio of 62:73 between rg1-1/rg1- 1;RG2/rg2-3 and rg1-1/rg1-1;RG2/RG2 is close to a 1:1 ratio [χ²(1, N = 135) = 0.9; 0.5 >p >0.1 for the probability that a deviation from a 1:1 ratio is due to chance]. The segregation ratio of 11:19 between RG1/rg1-1;rg2-3/rg2-3 and RG1/RG1;rg2-3/rg2-3 is somewhat skewed towards the RG1/RG1;rg2-3/rg2-3 genotype, but the chi-square analysis still suggest acceptance of the null hypothesis that 11:19 is not significantly different from 15:15 [χ²(1, N = 30) = 2.12; 0.5 >p >0.1]. Given that the latter data set is rather small, we discerned from this combined analysis a possible gametophytic lethality of the rg1-1;rg2-3 haplotype.

Dissection of siliques from rg1-1/rg1-1;RG2/rg2-3 and RG1/rg1-1;rg2-3/rg2-3 parent plants showed a reduced number of developed seeds and the presence of ~50% shrunken, white structures indicative of aborted ovules (Figure 1.3, Table 1.4). Silique length was not drastically affected in these semi-sterile heterozygotes compared to the single mutants, and no embryo lethals/seed abortions or seedling lethals were observed. Together, the

13 data suggest female gametophytic lethality of the combination of the two RanGAP null alleles.

1.4.3 Transmission of the rg1-1 allele in the rg2-3 background is normal through the male parent but completely blocked through the female parent

Analysis of the progeny of reciprocal crosses between RG1/rg1-1;rg2-3/rg2-3 X RG1/RG1;rg2-3/rg2-3 indicated that the rg1-1;rg2-3 genotype was fully transmissible through the male, but was 100% blocked through the female line (Table 1.5). This indicates lethality of rg1-1;rg2-3 female gametophytes. In addition, no morphological defects were observed in pollen grains from RG1/rg1-1;rg2-3/rg2-3 and rg1-1/rg1- 1;RG2/rg2-3 parents, supporting the notion that rg1-1;rg2-3 is male gametophyte viable. The fact that RG1 expression remained high while RG2 showed low expression in pollen according to GENEVESTIGATOR data (http;//www.genevestigator.ethz.ch; Zimmermann et al., 2004) encouraged the use of parental plants containing the RG1 wild-type allele instead of RG2 for the reciprocal crosses. Pollen tube transmission analysis in naturally selfed and aniline blue-stained RG1/rg1-1;rg2-3/rg2-3 and rg1- 1/rg1-1;RG2/rg2-3 pistils showed functional pollen and unfertilized mutant embryo sacs, indicating that the rg1-1;rg2-3 ovules are developmentally arrested prior to fertilization (Figure 1.4). The size and morphology of the arrested ovules appeared uniform and correlated with that of ovules of early mitotic stages. Together, these data suggest that rg1-1;rg2-3 female gametophytes are not viable, while rg1-1;rg2-3 pollen are viable and functional.

1.4.4 rg1-1;rg2-3 ovules arrest during early megagametogenesis

Microscopic floral observations and DIC optics of cleared ovules showed that the correlation between floral stages and ovule development described for Landsberg erecta (Ler) ecotype (Smyth et al., 1990; Schneitz et al., 1995) as shown in Table 1.2 was true

14 for WT Col (Figure 1.5) and WT Ws as well (Data not shown). Therefore, the same staging of floral and ovule development as described for Ler ecotype could be applied for WT Col and WT Ws. Subsequently, ovule development of rg1-1/rg1-1;RG2/rg2-3 parent plants was investigated (Figure 1.6 and Table 1.6). At stage 12-early, the majority of observed ovules were at the one- to two-nuclei stage without a vacuole and no difference between rg1-1;RG2 and rg1-1;rg2-3 ovules was observed. However, at stage 13-early, about half of the ovules had developed to an eight-nuclei or seven-nuclei stage, while the remaining half was distributed between one- to two-nuclei stage and more predominantly two-nuclei stage with vacuole. At stage 14-early, with half of the ovules at eight-nuclei or seven-nuclei stage, half of the arrested ovules were showing signs of degeneration. The DIC images in Figure 1.6 illustrate examples of a stage 12-early ovule, two ovules arrested with two-nuclei at stages 13-early and 14-early (presumed rg1-1;rg2-3 genotype) and one properly developed ovule at stage 13-early (presumed rg1-1;RG2 genotype).

These data suggest that the rg1-1;rg2-3 gametophytes were arrested predominantly after the first mitotic division following meiosis and degenerated subsequently. Similar results were obtained when analyzing RG1/rg1-1;rg2-3/rg2-3 parents (data not shown). Degeneration of arrested ovules has been reported for several gametophyte-lethal mutants (e.g. Pischke et al., 2002 and references therein). However, the mechanism that leads to the degeneration after developmental arrest is currently unknown. The arrests were evidently at the interphase as indicated by intact nuclei with visible nucleoli. The similar stage of arrest of the mutant ovules of both parents, RG1/rg1-1;rg2-3/rg2-3 and rg1- 1/rg1-1;RG2/rg2-3 suggests no differential effects by maternal (i.e. sporophytic mother plant) contribution of RG1 or RG2 towards ovule development and similar functionality for the two proteins in the underlying mechanisms in embryo sac development. Together, the observed arrest in early megagametogenesis, the absence of discernable embryo lethality/seed abortions and the absence of double mutant seedlings in repeated crosses, suggest that the RanGAP-deficient double mutants are lethal during the early mitotic phase of megagametophyte development.

15 1.4.5 RanGAPs are expressed in the developing embryo sac and pollen

Analysis of publicly available microarray data indicated the presence of RG1 mRNA in pollen, and both RG1 and RG2 mRNA in ovules (GENEVESTIGATOR data at www.genevestigator.ethz.ch; Zimmermann et al., 2004). Further, RG1- and RG2- promoter-driven GUS expression was observed in the developing and mature embryo sacs in the transgenic lines (Figure 1.7), thereby correlating RanGAP expression with a possible function(s) in the embryo sac (Transgenic lines were developed by Wang, H-J.). Figures 1.7A and B show β-Glucuronidase activity in stage 12-early ovule and embryo sac of a transgenic RG1-GUS line. A stronger staining was observed in the embryo sac than in the surrounding sporophytic tissue (Figure 1.7B), consistent with the interpretation that the RG1 promoter is active in the gametophyte. GUS expression detected in the integuments of ovules supports the possibility that RanGAP may be supplied from sporophytic tissues to developing embryo sacs. Strong GUS staining was also observed in pollen of the RG1-GUS plant, consistent with the published microarray data (Figure 1.7C). In the vegetative plant, mitotically active shoot and root meristems showed high GUS expression levels compared to a basal level of ubiquitous expression in the rest of the plant body (GUS expression analysis in the vegetative tissues was conducted by Wang, H-J.).

1.4.6 rg1-1/rg1-1;rg2-3/rg2-3 RanGAP knockout double mutants were rescued by complementation with RanGAP1 and RanGAP2 genomic constructs

To confirm RanGAP as the cause of lethality, rg1-1/rg1-1;RG2/rg2-3 plants were transformed with a native promoter-driven RG1 construct (Materials and Methods, Figure 1.8A) or a native promoter-driven RG2 construct (Materials and Methods, Figure 1.8B). Following transformation, rg1-1;rg2-3 double mutants were rescued in the T2 generation for both constructs as confirmed by genotyping. Figure 1.9A shows an example of a genotyping result. As controls, genotyping of a rg1-1/rg1-1;RG2/RG2 line

16 (lane 2) and a RG1/RG1;rg2-3/rg2-3 line (lane 3) are shown, confirming the identity of the respective allele-specific PCR products. Lane 1 shows a rescued line carrying the RG1-promoter RG1 transgene. Figure 1.9B shows an immunoblot using the RanGAP1- specific antibody against protein extracts from the same three lines. The viable rg1-1/rg1- 1;rg2-3/rg2-3 RG1 promoter-RG1 line presented shows more than twice as much RanGAP1 as the line homozygous for the wild-type RG1 allele and was phenotypically normal (Viable lines were also recovered that expressed ~25% of WT RG1). These data confirm that the female gametophyte lethality reported here is due to the combination of the two RanGAP null mutations, and not an independent event in the genetic background. This also supports redundancy between the two paralogs, RG1 and RG2, for female gametophyte development.

1.5 Discussion

1.5.1 RanGAP deficiency and lethality

In all organisms investigated, the complete loss of RanGAP is lethal. Mutants of the Saccharomyces cerevisiae (budding yeast) and Schizosaccharomyces pombe (fission yeast) homologs, rna1-1 and Sprnats respectively, are lethal at the restrictive temperatures (Hartwell, 1967; Kusano et al., 2004). The mutant of Fug1, the RanGAP homolog in mouse, is arrested at an early embryonic stage (DeGregori et al., 1994), while the RanGAP-RNAi mutants of Caenorhabditis elegans are lethal at embryonic or larval stages (Bamba et al., 2002). The data presented here appear the first to demonstrate that RanGAP is essential during gametogenesis. In addition, it was shown that Arabidopsis RanGAP1 and RanGAP2 are fully redundant at that stage. Surprisingly, RanGAP1 and RanGAP2 are essential for female, but not for male gametogenesis in the heterozygous plants studied here.

17 According to the known functions for RanGAP in other systems, the female gametophytes may undergo arrest due to malfunction of mitosis, defective nucleocytoplamic transport or perhaps a combination of both. In S. cereviceae rna1-1 mutant, deficiency in nuclear import of proteins was observed at the restrictive temperatures (Traglia et al., 1996). However, in the case of a series of temperature sensitive alleles of Sprnats, some showed no defects in nucleocytoplasmic protein transport but cell division was aberrant (Kusano et al., 2004). This suggests that different mutant alleles can reveal different RanGAP functions and that nucleocytoplasmic transport and mitotic functions might show different sensitivities to a reduction in the overall level of RanGAP.

XPO1/CRM1, the main exportin in the Ran cycle for proteins and required for chromosome segregation in mitosis, also showed female gametophytic lethality predominantly at the two-nuclear mitotic stage (Blanvillain et al., 2008). Thus, another component of the Ran cycle has shown to cause arrest in female gametophyte development. Further studies are necessary to dissect the underlying cause for arrest of RanGAP-deficient female gametophytes. It may be possible that a cause independent of the Ran cycle, nucleocytoplamic transport or mitosis was responsible for these arrests. Particularly because female gametophytes appear highly sensitive to both internal and external factors (Drews and Yadegari, 2002; Sun et al., 2004; Brukhin et al., 2005; Pagnussat et al., 2005), intrinsic stresses brought on by perturbed cellular processes may lead to lethality coincidently at a mitotic stage.

It is intriguing that the majority of mutant embryo sacs were able to survive up to two- nuclear mitotic stage. Parental contribution of RanGAP to the developing embryo sac could be a possible reason. Transport of mRNA or protein from the parent plant via cellular contact and/or cytoplasmic contribution from megaspore/microspore mother cell to the developing gametophyte has been speculated for other mutants that are deficient in an essential protein but are able to survive haploid development wholly or partially

18 (Drews et al., 1998; Kwee and Sundaresan, 2003; Brukhin et al., 2005; Shi et al., 2005; Blanvillain et al., 2008). In the case of RanGAP, the occurrence of parental contribution was indicated in a parallel experiment where rg1-1/rg1-1;RG2/rg2-2 and RG1/rg1-1;rg2- 2/rg2-2 mother plants produced full siliques lacking aborted ovules (i.e. rg1-1;rg2-2 ovules were not aborted), while rg1-1/rg1-1;rg2-2/rg2-2 mother plants which contain less RanGAP produced aborted rg1-1;rg2-2 ovules. Similarly, mother plant contribution of XPO1/CRM1 (another member of the Ran cycle and involved in nucleocytoplasmic transport and mitosis as described above), was shown to be required for gametophyte development (Blanvillain et al., 2008). It is reported that the cell destined to become the megaspore receives more than a fourth of the cytoplasmic content from the megaspore mother cell before meiotic division (Willemse and van Went, 1984) which may be a strategy in plant evolution to increase the supply of sporophytic material to the developing gametophyte. The fact that the mutant embryo sacs that lack a wild-type RanGAP allele aborted during development, while the embryo sacs containing a wild- type RanGAP allele developed normally, shows that the contribution of RanGAP from the parental heterozygotes (i.e. RG1/rg1-1;rg2-3/rg2-3 and, rg1-1/rg1-1;RG2/rg2-3) was not sufficient to support complete development of the rg1-1;rg2-3 female gametophyte.

Although two-nuclear mitotic stage arrests were predominant, a considerable number of mutant gametophytes also arrested at other stages including one- and four-nuclear mitotic stages and occasionally at megaspore mother cell stage and meiotic stages as well. Variable expressivity of the phenotypes is reportedly common in gametophyte mutants (Moore, 2002). Due to the haploid nature of the gametophytes, it is proposed that they are more susceptible to genetic and environmental modifiers (Brukhin et al., 2005). Alternately, the variability in stages of arrest could be due to a dilution effect from different initial RanGAP activities in the megaspore mother cell (Blanvillain et al., 2008). In a parallel experiment, the gametophytes developing on a further RanGAP knockdown plant rg1-1/rg1-1;rg2-2/rg2-2 showed much more variation in the stages of arrest ranging

19 from megaspore mother cell to post-mitosis, while some embryo sacs remained viable and fertilized (Chapter 2: Figure 2.3).

The percent ovule lethality although could be approximated to ~50% at large, variability ranging ~35%-60% per given silique of the heterozygote parents (RG1/rg1-1;rg2-3/rg2-3 and rg1-1/rg1-1;RG2/rg2-3) was encountered. Although higher percentage of abortion could be attributed to stress-induced lethality common in ovule development (Sun et al., 2004), lower percentages of aborted ovules may not be explained by a known mechanism for the RanGAP mutant other than ‘chance’. With statistical analyses using increased sample sizes, if a tendency for lower aborted rg1-1;rg2-3 ovules could be established, it would constitute a recapitulation of a scenario similar to the NOMEGA mutant defective in the functioning of the anaphase-promoting complex/cyclosome which showed no more than 30% female gametophytic abortion (Kwee and Sundaresan, 2003).

Similar to NOMEGA mutant, embryo lethality was not observed in the heterozygous RanGAP mutant and nor were any RanGAP double mutant seedlings recovered despite extensive screening. In addition, aniline blue stained pistils showed that the fully developed ovules were each delivered with a pollen tube thereby eliminating the possibility that a considerable number of mutant ovules may have arrested post- gametophyte development and pre-fertilization. These results suggest that the observed arrested ovules constitute the majority of, if not all, embryo sacs containing the mutant genotype. Therefore, it is interesting to consider the possibility that an unfavorable bias may lead to the development of a less number of mutant ovules in the ovary of the heterozygotes. If complete randomness in megaspore generation from the meiotic nuclei was assumed, such reduction in the percentage mutant ovules is improbable. Therefore, it remains open whether there is a preferential selection for wild-type meiotic products to becoming the functional megaspore. As described above, non-equal distribution of the cytoplasm in favor of the future megaspore has been reported (Willemse and van Went, 1984), which supports a favorable bias towards the functional megaspore.

20 1.5.2 RanGAP1 and RanGAP2 are functionally redundant in female gametophyte development

The fact that both RG1;rg2-3 and rg1-1;RG2 ovules developed normally on the heterozygous parents and the mutant rg1-1;rg2-3 ovules could be rescued by both RG1 and RG2 transgenic constructs independently shows functional redundancy of RanGAP1 and RanGAP2 in female gametophyte development. This is in contrast to the RanGAP2- specific function reported for disease resistance against PVX (Sacco et al., 2007; Tameling and Baulcombe, 2007), but is in accordance with the proposed cell plate establishment function performed redundantly by the two plant RanGAPs (Xu et al., 2008). This redundancy of plant RanGAPs may have hampered identification of their role in embryo sac development by numerous forward genetic screens (Feldmann et al., 1997; Moore et al., 1997; Bonhomme et al., 1998; Christensen et al., 1998; Howden et al., 1998; Grini et al., 1999; Shimizu and Okada, 2000; Pagnussat et al., 2005). Therefore, a reverse genetic approach, as followed in this study, was vital to reveal the functional aspects of RanGAP in megagametophyte development.

1.5.3 Female gametophytes are arrested during a mitotic phase

The nearly ubiquitous pattern of 1) RanGAP mRNA accumulation (GENEVESTIGATOR data at www.genevestigator.ethz.ch; Zimmermann et al., 2004) and 2) promoter-reporter activity supports the notion that RanGAP encodes an essential protein. While the high expression in mitotic tissues suggests a mitotic function, the ubiquitous expression through out the plant body may suggest a mitosis-unrelated essential function, possibly nucleocytoplasmic transport. However, the high expression of RanGAP in the mitotic tissues may also indicate an increased demand for a housekeeping cellular process for which RanGAP is required, yet not directly related to mitosis. If embryo sac lethality did result due to a deficient mitotic function of RanGAP, the nuclear arrest-phenotype resembles the mutants of positive cell division regulators (Springer et al.,

21 1995; Springer et al., 2000; Capron et al., 2003; Kwee and Sundaresan, 2003; Wu et al., 2005; Binarova et al., 2006; Pastuglia et al., 2006; Barrero et al., 2007). On the contrary, mutants of negative regulators of cell division have reported supernumerary nuclei in the embryo sac as described in section 1.2. The exact underlying cause for the nuclear arrest in the RanGAP deficient embryo sacs; whether it is a deficiency in a direct mitotic function of RanGAP, a downstream effect of a deficient nucleocytoplasmic transport function or an independent unrelated function remains to be determined.

1.5.4 Embryo sacs are affected more than the pollen

The commonly observed phenomena in the gametophtytic mutants are general gametophytic lethality that affects both pollen and ovules, and a leaky nature of lethality that allows transmission of some percentage of the mutation to the next generation (Drews et al., 1998; Drews and Yadegari, 2002). However, RanGAP knockout mutation in the heterozygous parents showed total lethality and limited to the female gametophyte alone. Total lethality implies an absolute essentiality of RanGAP in female gametophyte development. Pollen from these mother plants, on the other hand, not only survived the full phase of development including the mitotic divisions but also functioned normally. How pollen that shares many developmental processes common with embryo sacs including mitosis, vacuole formation, cell expansion, subcellular migration, cellularization and cell wall formation (Drews et al., 1998) could be thus spared from effects of aberrance of an essential gene function is puzzling. Accordingly, the existence of a vital RanGAP-function that is specific to female gametophyte development alone can not be ruled out. Nevertheless, high RanGAP1 expression reported in GENEVESTIGATOR data for pollen was confirmed by promoter-GUS expression as well. Alternatively genetic redundancy that compensates for RanGAPs in microspores alone can not be ruled out (Shi et al., 2005). However, in a parallel experiment, further reduction of RanGAP in the parental tissue in trans-heterozygous plants (rg1-1/rg1- 1;rg2-2/rg2-3) showed a slight reduction in mutant pollen performance as well. When

22 WT Col pistils were pollinated with pollen from rg1-1/rg1-1;rg2-2/rg2-3 plants, the knockout rg2-3 allele showed only 77.5% genetic transmission through the pollen compared to the 100% genetic transmission rate of the competing knockdown rg2-2 allele (Chapter 2: Table 2.2) (Allelic/genetic transmission as described in Howden et al., 1998).

More severe effects in female gametophyte development compared to the male have been reported for mutants deficient in essential genes that are not directly involved in mitosis (Coury et al., 2007 and references therein). This may suggest that the differences between requirements for female and male gametophyte development could place a higher demand by the female gametophyte towards the functions of common essential genes such as genes involved in nucleocytoplasmic transport. The development process of the embryo sac involves more elaborate changes compared to pollen, including dramatic increase in size while no or little growth takes place in microspore development (Shi et al., 2005). Thus, less requirement of RanGAP for pollen development and function could be envisioned due to lower housekeeping requirements. On the other hand, more parental contribution of RanGAP in pollen compared to embryo sac may be possible and/or normal pollen performance may be aided by the transmittance of pollen tube through the tissue of pistil which may supplement the growing pollen with RanGAP due to close tissue contact.

With respect to direct effects on mitosis, one striking difference is the additional mitotic division cycle of the haploid nuclei in embryo sac development compared to pollen, thus requiring material to support an extra round of division in the embryo sac (Shi et al., 2005). Further, it could be possible that the small size or the simpler network of cellular processes may have enabled the pollen to survive through the divisions with the least expenditure of RanGAP. Similar to RanGAP, less pronounced male gametophytic defects have been shown in several mutants of genes involved in cell division such as γ-Tubulin (Pastuglia et al., 2006), NOMEGA mutant of Anaphase Promoting Complex/Cyclosome

23 (APC/C) (Kwee and Sundaresan, 2003) and mutant of chromatin-remodeling protein 11 (CHR11) (Huanca-Mamani et al., 2005). SLOW WALKER1 is a mutation in a WD40 protein that is essential only for female gametogenesis, however in this case a redundant function in the male gametophyte cannot be excluded (Shi et al., 2005). Interestingly, a double null mutant of the two genes encoding the Arabidopsis karyopherin XPO1/CRM1, the main exportin for proteins, also showed predominantly female gametophytic lethality (Blanvillain et al., 2008). Similar to the data described here for RanGAP mutants, the embryo sac arrests in the XPO1/CRM1 mutant was predominantly at the two-nuclear mitotic stage. In addition to its function in Ran-dependent nuclear export, XPO1/CRM1 is also involved in the mitotic functions of the Ran cycle in animals (Arnaoutov and Dasso, 2005; Arnaoutov et al., 2005). Thus, with respect to gene function (i.e. Ran cycle and mitosis) and phenotype (more prevalent aberrance in female than male gametophyte), the two mutants XPO1/CRM1 and RanGAP show striking overlap suggestive of a perturbed Ran cycle function involved in mitosis.

1.5.5 Outcomes of the study and future prospects

This work contributes to the understanding of function of RanGAPs in the haploid phase of Arabidopsis development. Given the redundancy revealed for the RanGAP paralogs in embryo sac development, functional dissection herein may not have been possible by a classical forward genetic approach. The mitotic arrest of the mutant female gametophytes, which could be possible via a direct mitotic effect or a downstream effect of an essential function such as nucleocytoplamic transport mediated by RanGAP, encourages a hypothesis that RanGAP’s classical functions may be conserved in planta as well.

In order to evaluate whether the arrest of female gametophytes occur as a secondary downstream effect of perturbed nucleocytoplasmic transport due to RanGAP deficiency, a transport assay could possibly be devised. Methodology including possible marker proteins could be adopted from the available literature (Sanderfoot et al., 1996; Haasen et

24 al., 1999; Ward and Lazarowitz, 1999; Kusano et al., 2004; Cheng et al., 2009; Hong et al., 2011). Due to the internal nature of the female gametophyte and the challenges in isolating embryo sacs as live protoplasts for real time assays, confocal microscopy of fluorescence-tagged nucleocytoplasmic shuttling marker proteins expressed using a female gametophyte-specific promoter (such as pFM1 used in Huanca-Mamani et al., 2005) or histological sectioning of ovules and immunodetection of shuttling marker proteins may be required.

Microarray experiments on isolated two-nuclear embryo sacs of arrested rg1-1;rg2-3 ovules (12 ‘late’ stage) may provide valuable insights on the cause of arrest, particularly to evaluate whether defective mitosis is causal for the phenotype. Expression of cell cycle related genes could be thus evaluated in comparison to a suitable control. However, due to the complications such as the mixed Col/Ws background in the mother plant and possible parental transport of gene expression products to the embryo sac (Drews et al., 1998; Kwee and Sundaresan, 2003), multiple controls may need to be used for the comparisons. Possible controls include total two-nuclear embryo sacs of the heterozygous mother plants (two-nuclear rg1-1;rg2-3 and heterozygous rg1-1;RG2 or RG1;rg2-3 ovules from 12 ‘mid’ stage), seven-nuclear heterozygous rg1-1;RG2 or RG1;rg2-3 embryo sacs isolated from 12 ‘late’ stage of the heterozygous mother plants and two- nuclear embryo sacs of the single mutant parent (RG1/RG1;rg2-3/rg2-3 or rg1-1/rg1- 1;RG2/RG2) that co-segregate with the heterozygous mother plant. A practically less challenging microarray approach may include a subtractive microarray (Jones-Rhoades et al., 2007) involving arrested whole ovules containing two-nuclear rg1-1;rg2-3 embryo sacs from the heterozygote and arrested whole ovules of determinate infertile 1 (dif1) mutant lacking an embryo sac (Jones-Rhoades et al., 2007). dif1 mutation does not affect the sporophyte development including the integuments, which is advantageous to serve as a control. However, the dif1 mutant (SALK_091193, Columbia background) would firstly need to be developed in a Col/Ws mixed background similar to the heterozygote by crossing the dif1 mutant (Col) with Ws wild-type.

25 In Chapter 2, it is suggested that development of vegetative meristems in RanGAP- deficient rg1-1/rg1-1;rg2-2/rg2-3 plants may be perturbed due to a defective cell cycle commitment, likely in G1/S phase transition. In order to evaluate whether ovules of RanGAP-deficient mutants arrest as a result of defective G1/S phase transition, rescue experiments with mutations in negative regulators downstream in this pathway could be attempted. The developmental sensitivity of embryo sacs lead to their lethality due to a myriad of causes (Drews and Yadegari, 2002; Sun et al., 2004; Brukhin et al., 2005; Pagnussat et al., 2005). Therefore, pursue of an embryo sac-arrest phenotype via genetic crosses with other mutants is challenging as they may arrest as a result of compiled cellular stress effects instead of the bonafide molecular cause.

Therefore, performing rescue experiments instead appear a feasible approach. Retinoblastoma protein is a negative regulator of cell cycle commitment acting downstream of Cyclin D induction in G1/S transition and represses the E2F transcription factors that initiate cell division functions in the S phase (den Boer and Murray, 2000; Ebel et al., 2004). The phosphrylation cascade triggered by Cyclin D induction is shown to release retinoblastoma protein from inhibitory binding to E2F transcription factors, upon release of which the E2F transcription factors are able to facilitate transcription of the S-phase genes (den Boer and Murray, 2000). Mutations in retinoblastoma homolog in Arabidopsis [i.e. retinoblastoma-related (RBR1) protein] have shown to result in supernumerary nuclei in female gametophytes (Ebel et al., 2004), presumably due to uncontrolled nuclear division. Experiments in Chapter 2 suggested a possibility that RanGAP could act upstream of Cyclin D induction in G1/S transition of cell division in the vegetative meristems. Therefore, if mutations in RBR1 are able to rescue the female gametophyte arrests observed in rg1-1;rg2-3 ovules, it would provide valuable insight to RanGAP’s involvement in G1/S transition of mitotic division in female gametophyte development. It would be intriguing to perform this genetic rescue experiment on rg1- 1;rg2-3 female gametophytes developing on the heterozygotes rg1-1/rg1-1;RG2/rg2-3 and RG1/rg1-1;rg2-3/rg2-3. However, due to the complete sterility of female

26 gametophytes containing rg1-1;rg2-3 and rbr1 mutant alleles independently (Ebel et al., 2004), this approach is not feasible in the heterozygotes. Therefore, rescue of rg1-1;rg2-2 ovules developing on the knockdown mutant rg1-1/rg1-1;rg2-2/rg2-2 could be attempted because a percentage of rg1-1;rg2-2 embryo sacs would be viable due to the leaky nature of the arrest phenotype. Genetic crosses between the rg1-1/rg1-1;rg2-2/rg2-2 mother plant and rbr1 mutants would potentially enable obtaining mutant embryo sacs containing the rg1-1;rg2-2;rbr1 genotype in order to evaluate phenotypic rescue. If the underlying cause for arrests of embryo sacs developing on heterozygotes and the rg1- 1/rg1-1;rg2-2/rg2-2 mutant is similar, a positive rescue result would provide valuable insights.

To evaluate if defective Cyclin D induction in G1/S transition is a cause of arrest of rg1- 1;rg2-3 female gametophytes developing on the heterozygotes, in situ hybridization experiments to quantify Cyclin D transcript levels in mutant versus heterozygous rg1- 1;RG2 or RG1;rg2-3 ovules at two-nuclear embryo sac stage could be performed. Alternatively, rescue experiments of rg1-1;rg2-3 female gametophytes via Agrobacterium-mediated transformation with Cyclin D3 or Cyclin D2 expression constructs could be attempted. A female gametophyte-specific promoter (such as pFM1 used in Huanca-Mamani et al., 2005) or a native Cyclin D promoter would be required for rescue due to possible complications associated with using the CaMV35S promoter for gene expression in female gametophytes (Jenik and Irish, 2000; Huanca-Mamani et al., 2005).

27 A 200bp rg1-1

ATG

Promoter Ex In Ex

5’ UTR ORF 3’ UTR

B rg2-3

ATG

Promoter Ex In Ex

5’ UTR ORF 3’ UTR

Figure 1.1: Schematic representation of the gene structure and T-DNA insertions in RanGAP1 and RanGAP2. (A) RanGAP1. (B) RanGAP2. ORF, open reading frame; Ex, Exon; In, Intron; ATG, open reading frame start position; blocked arrow, open reading frame end position. Triangles depict the positions of T-DNA insertions. Scale bar: 200 bp.

28 A

WT rg1-1

70 kD α -RanGAP1

Coomassie

WT (Ws) rg2-3

70 kD α - RanGAP2

Coomassie

Figure 1.2: Immunoblot analysis confirms rg1-1 and rg2-3 as knockout (null mutant) lines. (A) rg1-1 and (B) rg2-3 alleles lack observable protein expression of RanGAP1 and RanGAP2 respectively. Total protein extracts from (A) 10 day-old and (B) 8 day-old Arabidopsis seedlings containing each mutant allele in homozygous condition were incubated with (A) anti-RanGAP1 and (B) anti-RanGAP2 antibodies. WT, Columbia wild-type; WT (Ws), Wassilkja wild-type. Blot (A) was generated by Xu, X.M. (Xu, 2007). Coomassie brilliant blue staining of replica gels is shown at the bottom as the loading control.

Figure 1.3: Siliques of heterozygous plants are semi-sterile. (A) Dissected silique from rg1-1/rg1-1;RG2/rg2-3 parent containing a reduced number of seeds. Arrows point to the undeveloped ovules which appear as shrunken white masses. (B) Part of a silique from a wild-type parent showing full seed set. Scale bars: 100 µm.

Figure 1.4: RanGAP mutants are pre-fertilization lethal. Fluorescence micrographs showing pollen tube transmission in naturally selfed and aniline blue-stained pistils from (A) rg1-1/rg1-1;RG2/rg2-3 heterozygote and (B) wild-type control. Arrow, fertilized rg1-1;RG2 ovule; Arrowhead, aborted rg1-1;rg2-3 ovule. Note that mutant ovules in (A) do not attract pollen tubes and assume smaller kidney shaped morphology which is characteristic of ovules containing female gametophytes of early mitotic stages. Pollen tubes are seen entering the developing seeds in (A). Wild-type (B) contains uniformly- sized seeds which are supplied with pollen tubes. Scale bars: 100 µm.

31 Figure 1.4

A B

32 10 11 11 12‐III 12‐IV

12‐I

12‐V 12‐VI 33 12‐I 12‐II

Figure 1.5: Floral, ovule and embryo sac development stages of Columbia correlate with those described for Landsberg erecta. Optical sections of cleared ovules from a wild-type (Col) parent observed under DIC microscopy. Numeric denotations indicate stage of floral development as described in Table 1.2. Red arrows point to the nuclei; green arrow points to the vacuole. Scale bars: 25 µm. A 12‐early B 13‐early

rg1‐1; RG2 or rg1‐1; rg2‐3 rg1‐1; rg2‐3 C 14‐early D 13‐early

rg1‐1; rg2‐3

Fgure 1.6: The majority of RanGAP mutant ovules are aborted at the two-nuclear embryo sac stage. Optical sections of cleared ovules from a rg1-1/rg1-1;RG2/rg2-3 parent observed under DIC microscopy. (A) Developing rg1-1;RG2 or rg1-1;rg2-3 ovule from floral stage 12-early containing two nuclei and prior to vacuole formation. (B) Arrested rg1-1;rg2-3 ovule from floral stage 13-early containing two nuclei separated by a vacuole. (C) Degenerating rg1-1;rg2-3 ovule from floral stage 14-early containing two nuclei separated by a vacuole. (D) Developing rg1-1;RG2 ovule from floral stage 13- early. Floral and ovule development stages are as described by Smyth et al., 1990 and Schneitz et al., 1995. ‘early’ refers to the beginning of the respective floral stage. Arrows point to the nuclei; white dotted lines encircle the position of nuclei; P, unfused polar nuclei; E, egg cell nucleus; S, synergid nuclei. Scale bar: 50 µm.

34 A C

Stage 12 early

Figure 1.7: RanGAP promoter-GUS activity is detected in embryo sacs and pollen. GUS-staining of (A) ovule (B) embryo sac and (C) flower with pollen of a RG1 promoter-GUS transgenic line. Arrow in (B) indicates the position of embryo sac, which is more intensely stained compared to the surrounding sporophytic tissue of the ovule. (C) was generated by Wang, H-J. Scale bars: 50 µm.

35 A

ATG Stop

~0.73kb RG1 promoter 5’UTR Intron 5’UTR Coding region 3’UTR IR GUS

RG1 promoter used for GUS assay Cloned-in ~2.2kb fragment

ATG Stop

~1.2kb RG2 promoter 5’UTR Intron 5’UTR Coding region 3’UTR

Cloned-in ~4kb fragment

Figure 1.8: Schematic representation of the genomic constructs used for complementation. (A) Native RG1- and (B) native RG2-promoter driven constructs. Yellow regions in (A) depict elements of the RG1-promoter-GUS construct and the red region represents the newly introduced ~2.2 kb genomic region of RG1. IR, intergenic region of ~300 bp included in the ~2.2 kb genomic fragment. ‘ATG’ and ‘Stop’ refer to start and end of the coding region respectively. Schematics are not drawn to scale.

1 G R ; ; A 1 + 1 ; - - 1 3 1 -3 1 - g 2 g 2 G 2 /r g /r R g r G / /r -1 / -1 R 1 1 3 1 / G 3 g - 2 - r 2 rg R 2 g G g r R r

rg1-1

RG2

rg2-3

RG1 or +RG1

+RG1 RG1

rg1-1/rg1-1; rg1-1/rg1-1; RG1/RG1; rg2-3 /rg2-3 RG2/RG2 rg2-3 /rg2-3 + RG1

α-RanGAP1

Coomassie

Figure 1.9: Mutant lethality is rescued by genomic RanGAP constructs. (A) Genotyping PCR result of a rescued mutant plant in the T2 generation resulting from transformation of rg1-1/rg1-1;RG2/rg2-3 with the native RG1 construct. As controls for the PCR reactions, genotyping of rg1-1/rg1-1;RG2/RG2 and RG1/RG1;rg2-3/rg2-3 plants were performed. +RG1, transgenic RanGAP1; RG1, endogenous RanGAP1. (B) Immunoblots of the same plants with anti-RanGAP1 antibody. Rosette leaves of 21 day- old plants were used. Coomassie brilliant blue staining of a replica gel is shown at the bottom as the loading controls.

37 Table 1.1: Primers used in this study

38 Table 1.2: Floral stages, landmark characteristics and female gametophyte development (Adapted from Smyth et al., 1990 and Schneitz et al., 1995)

39 Table 1.3: Expected and observed F3 segregation ratios derived from a cross between rg1-1/rg1-1;RG2/RG2 and RG1/RG1;rg2-3/rg2-3

F2 parent: rg1-1/rg1-1;RG2/rg2-3 F2 parent: RG1/rg1-1;rg2-3/rg 2-3 Progeny Expected Observed1 Progeny Expected Observed2 (%) (%) (%) (%)

rg1-1/rg1-1; 67.5/135 62/135 RG1/rg1-1; 15/30 11/30 RG2/rg2-3 (50) (46) rg2-3/rg2-3 (50) (37)

rg1-1/rg1-1; 33.8/135 73/135 RG1/RG1; 7.5/30 19/30 RG2/RG2 (25) (54) rg2-3/rg2-3 (25) (63)

rg1-1/rg1-1; 33.8/135 0/135 rg1-1/rg1-1; 7.5/30 0/30 rg2-3/rg2-3 (25) (0) rg2-3/rg2-3 (25) (0)

F2 RG1/rg1-1;rg2-3/rg 2-3 and rg1-1/rg1-1;RG2/rg2-3 plants were self-fertilized and F3 segregation ratios were determined. ‘Expected’ refers to outcome expected according to Mendelian genetics. ‘Observed’ refers to outcome obtained.

1 χ²(1, N = 135) = 0.9; 0.5 >p >0.1. 2 χ²(1, N = 30) = 2.12; 0.5 >p >0.1.

N refers to the number of individuals genotyped.

40 Table 1.4: Number of aborted ovules from rg1-1/rg1-1;RG2/rg2-3 and wild-type (Col) siliques

Longest siliques from three well-grown plants were used for counts from each parent. ‘Aborted ovules’ refer to miniscule white masses. ‘Aborted seeds’ refer to partially developed smaller seeds. ‘Total’ refers to the sum of developed seeds, aborted ovules and aborted seeds.

41 Table 1.5: Transmission of the rg1-1;rg2-3 genotype through the male and female gametophytes

Cross RG1/RG1; RG1/rg1-1; Number of TE TE rg2-3/rg2-3 rg2-3/rg2-3 progeny Female Male Female Male genotyped % %

RG1/rg1-1; RG1/RG1; 62 0 62 0 NA rg2-3/rg2-3 rg2-3/rg2-3

RG1/RG1; RG1/rg1-1; 29 32 61 NA 108 rg2-3/rg2-3 rg2-3/rg2-3

TE, transmission efficiency; NA, not applicable. TE was calculated according to Howden et al. (1998). TE= progeny containing rg1-1;rg2-3 /progeny lacking rg1-1;rg2-3 X 100%.

42 Table 1.6: Percentage of female gametophytes at selected stages of development in the siliques of semi-sterile heterozygote rg1-1/rg1-1;RG2/rg2-3

Floral Stage of female gametophyte development Total number stage Percent ovules of ovules analyzed 1 nu. and 2 2 nu. 8 nu. Degenerated Other nu. without with vac. and vac. 7nu.

12 87.0 8.2 - - 4.8 62 early

13 6.5 36.4* 49.4 - 7.8 77 early

14 - 24.3 45.7 25.7 4.3 70 early

Ovules with unclear gametophyte development stages were not counted. ‘early’ refers to the beginning of the floral stage showing hallmark features of the corresponding stage. ‘14 early’ refers to the floral developmental stage that is 13 h after emasculation at ‘l3 early’ in order to prevent fertilization. nu., nuclei; vac., vacuole. *Some gametophytes already degenerating at this stage are included.

43 CHAPTER 2

REQUIREMENT OF RANGAP FOR VEGETATIVE DEVELOPMENT AND CELL DIVISION IN ARABIDOPSIS THALIANA

44 2.1 Abstract

RanGAP, the GTPase-activating protein of the small GTPase Ran, provides spatial information for nucleocytoplasmic transport via the Ran cycle and various mitotic processes in yeast and animals. Arabidopsis thaliana contains two paralogous copies of RanGAP, namely RanGAP1 and RanGAP2. In an attempt to investigate the functions of plant RanGAPs on a whole-organism level, T-DNA insertion mutants were analyzed. In the absence of observable vegetative phenotypes in the single knockout mutants and female-gametophyte lethality in the double knockout mutants, knockdown mutants were constructed and analyzed. A seedling phenotype with a mild delay in shoot and root development was observed in a knockdown mutant; rg1-1/rg1-1;rg2-2/rg2-2, which was more prominently manifested in a trans-heterozygous mutant; rg1-1/rg1-1;rg2-2/rg2-3 carrying a further reduced level of RanGAP. In rg1-1/rg1-1;rg2-2/rg2-3 plants, root growth was severely retarded and the disrupted shoot development from the primary shoot apical meristem (SAM) was followed by growth arrest of the primary shoot and multiple shoot initiation from adventitious primordia. Seedling death was observed in the most severely affected seedlings. Adult plants that developed from the multiple adventitious shoots were bushy and stunted. Complementation with AtRanGAP1 or AtRanGAP2 overexpression constructs rescued these phenotypes independently.

The observed mutant seedling phenotypes showed similarity to reported cell cycle mutants that were defective in genes functioning in different phases of the cell cycle. Further, these phenotypes were similar to wild-type plants treated with chemicals or radiation that perturb different phases of the cell cycle. Accordingly, the shoot and root apex sizes were reduced in rg1-1/rg1-1;rg2-2/rg2-3 and cell division was restricted to compact areas in the primary vegetative apices. Supplementation with sucrose, which was shown to promote cell cycle re-entry at G1/S boundary in cell suspension and root tip cultures or G2/M transistion in an Arabidopsis mutant, rescued both root and shoot vegetative phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 plants. Partial rescue to almost

45 complete rescue were observed depending upon the environmental conditions. Concomitantly, mitotic activity in the vegetative apices was restored, supporting defective cell division as the primary cause of the phenotypes in rg1-1/rg1-1;rg2-2/rg2-3 plants. Sucrose supplementation was able to suppress the development of adventitious shoots while promoting the growth and development of the primary SAM, thereby giving rise to adult plants with restored wild-type-like vegetative stature and height. Conversely, sucrose depletion resulted in lethality of rg1-1/rg1-1;rg2-2/rg2-3 prior to seedling establishment. Taken together, the results suggest that RanGAP is important for vegetative plant development via a cell division role and that the two Arabidopsis RanGAPs are redundant for this function. Sucrose rescue might suggest an involvement of RanGAP in cell cycle progression at G1/S or G2/M transitions.

46 2.2 Introduction

2.2.1 Ran cycle and the functions of RanGAP in animals and yeast

Ran (a small GTPase of the Ras superfamily) is essential for nucleocytoplamic transport via the Ran cycle in yeast and animals, and in animal mitosis (Dasso, 2002; Hetzer et al., 2002; Arnaoutov and Dasso, 2003; Quimby and Dasso, 2003; Arnaoutov and Dasso, 2005). The functioning of the Ran cycle requires switching of Ran between its GTP- and GDP-bound forms but the intrinsic rate of GDP hydrolysis by Ran per se is low (Bischoff et al., 1994; Bischoff and Ponstingl, 1991). The Ran GTPase-Activating Protein (RanGAP) stimulates the intrinsic GTPase activity of Ran which facilitates the hydrolysis of the Ran-bound GTP to GDP while the guanine nucleotide exchange factor (RanGEF/RCC1) replaces the Ran-bound GDP with a GTP molecule (Bischoff and Ponstingl, 1991; Bischoff et al., 1994; Bischoff et al., 1995b; Seewald et al., 2003).

RanGAP has been associated with essential functions in nucleocytoplasmic transport via the Ran cycle (Bischoff et al., 1995a; Traglia et al., 1996; Hutten et al., 2008) as well as mitosis in animals and yeast. Schizosaccharomyces pombe (fission yeast) RanGAP- mutants demonstrated an essential function for centromeric chromatin condensation and chromosome segregation, while nucleocytoplasmic transport was normal in the majority of them (Kusano et al., 2004). In animal mitosis, functions of RanGAP include spindle assembly, centrosome duplication, chromosome alignment, segregation and decondensation as well as nuclear envelope re-assembly presumably via Ran cycle functions (Hetzer et al., 2000; Zhang and Clarke, 2000; Gruss et al., 2001; Bamba et al., 2002; Zhang et al., 2002; Arnaoutov and Dasso, 2003; Arnaoutov and Dasso, 2005; Ciciarello et al., 2007). Given these essential functions of RanGAP, null mutants in yeast and animals are lethal (DeGregori et al., 1994; Traglia et al., 1996; Bamba et al., 2002; Kusano et al., 2004).

47 2.2.2 RanGAPs in Arabidopsis thaliana

Arabidopsis thaliana carries 2 paralogs of RanGAP (RG), namely RanGAP1 (RG1) and RanGAP2 (RG2), which share ca. 60% identity with each other and ca. 20% identity with either the Saccharomyces cereviceae ortholog (ScRna1p) or human RanGAP ortholog (HsRanGAP) (Rose and Meier, 2001). The GAP (i.e. GTPase-Activating Protein) activity is conserved in Arabidopsis RanGAPs as shown by complementation of the ScRna1p mutant, rna1-1 (Ach and Gruissem, 1994; Merkle et al., 1994; Pay et al., 2002). Specific subcellular localizations of RanGAP during the cell cycle determine the local RanGDP concentration which is thereby considered to provide spatial cues vital for functions of RanGAP in the Ran cycle (Joseph et al., 2002; Joseph et al., 2004). As in animal cells, nuclear envelope localization of RanGAP is conserved in differentiated cells and during interphase of mitotic cells in the root tip of Arabidopsis (Rose and Meier, 2001; Pay et al., 2002; Jeong et al., 2005). Similarly, mitotic localizations of RanGAP seen in animals such as kinetochores and spindle vicinity are conserved in plants as well (Matunis et al., 1998; Joseph et al., 2002; Pay et al., 2002; Xu et al., 2008). This suggests that plant RanGAPs may have similar nucleocytoplasmic transport and cell division functions as in animal cells.

In addition, Arabidopsis RanGAP1 shows plant-specific localizations throughout mitosis and cytokinesis, including the preprophase band, cortical division site, phragmoplast and the outwardly growing rim of the nascent cell plate (Jeong et al., 2005; Xu et al., 2008). These localizations of RanGAP1, together with misplaced cell walls and cell wall stubs in RanGAP-depleted inducible RNAi lines, have led to the hypothesis that RanGAP is involved in proper cell division plane establishment in plant cytokinesis via acting as a persistent positional marker of the future cell plate (Xu et al., 2008). Other known functions of plant RanGAPs are limited to the RanGAP2-specific involvement in disease resistance against potato virus X via an unknown, nuclear import-independent function (Sacco et al., 2007; Tameling and Baulcombe, 2007) and a redundant function for the two

48 RanGAPs in female gametophyte development in Arabidopsis (Rodrigo-Peiris et al., 2011).

Despite conserved localization pattern of animal and plant RanGAPs, their targeting mechanisms differ. In mammalian RanGAP, a unique C-terminal domain upon sumoylation binds to the nucleoporin RanBP2/Nup358 and colocalization of these two proteins together mediates targeting of RanGAP to nuclear envelope and mitotic localizations (Matunis et al., 1998; Joseph et al., 2002; Joseph et al., 2004). In Arabidopsis RanGAPs, the plant-specific N-terminal WPP domain mediates targeting to nuclear envelope and mitotic localizations (Rose and Meier, 2001; Jeong et al., 2005; Xu, 2007). Interaction between the WPP domain and nuclear envelope-anchored WIP and WIT proteins mediate RanGAP localization to the nuclear envelope (Xu et al., 2007a; Zhao et al., 2008).

2.2.3 Shoot and root development in Arabidopsis thaliana

The embryonic/primary shoot apical meristem (SAM) and the embryonic/primary root apical meristem (RAM) contain the major banks of cycling cells in a seedling that determine the apical-basal growth of the plant body (Sharma and Fletcher, 2002; Weigel and Jurgens, 2002; Baurle and Laux, 2003; Veit, 2004). The SAM consists of three zones in dicots, which are 1) the central zone at the apex that consists of slowly dividing stem cells, 2) the peripheral zone that encircles the central zone and contains faster dividing cells that give rise to leaf primordia, and 3) the rib zone placed below the central and peripheral zones that provides cells for the central tissues of the stem (Steeves and Sussex, 1989; Van Lijsebettens and Clarke, 1998; Nakajima and Benfey, 2002; Sharma and Fletcher, 2002; Jacqmard et al., 2003). The Arabidopsis root tip consists of 3 zones: 1) the meristematic zone/cell division zone/RAM which supplies a bulk of root cells for root growth and is surrounded by the root cap (Dolan et al., 1993; Schiefelbein and Benfey, 1994; Ubeda-Tomas et al., 2009), 2) the cell elongation zone which consists of rapidly

49 proliferating and expanding cells (Schiefelbein and Benfey, 1994; Siddiqui et al., 2003), and 3) the cell differentiation zone where the elongated cells differentiate for specialized functions (Dolan et al., 1993; Baluska et al., 1996).

Root growth/length is determined by cell division and elongation in the root tip which thereby relies on the activities in the meristematic and cell elongation zones (Dolan et al., 1993; Schiefelbein and Benfey, 1994; Siddiqui et al., 2003; Ubeda-Tomas et al., 2009). The rapid cell division that takes place in the meristematic zone/cell division zone/RAM during the first 4-6 days after germination establishes the size of the RAM which is vital for root growth (Ubeda-Tomas et al., 2009). Not only does it establish a bulk of dividing cells that determines the dimensions of the RAM and cell numbers of the root, it also supplies cells to undergo rapid elongation in the elongation zone (Ubeda-Tomas et al., 2009). The elongation zone that consists of rapidly dividing and elongating cells is also responsible for root growth via increasing cell numbers and anisotropic/longitudinal expansion of cells (Schiefelbein and Benfey, 1994; Siddiqui et al., 2003).

While the shape of cells differs demarcating the root meristem zone from the elongation zone (i.e. isodiametric cells in the meristem versus anisotropic/elongated cells in the elongation zone), the elongation zone is demarcated from the differentiation zone by the presence of root hairs which are characteristic of differentiated root cells. Thus, the distal (i.e. more apical) end of the differentiation zone bordering the cell elongation zone is marked by epidermal cells containing root hairs (Dolan et al., 1993). The meristematic zone consists of 4-7 mitotically quiescent stem cells at the core termed the ‘quiescent center’ (QC) which by occasional divisions give rise to the rapidly dividing stem cells called the ‘initial cells’ that surround the quiescent center (Nawy and Benfey, 2001). The central/columella root cap is established by the ordered division of a layer of the stem cells placed immediately distal to the quiescent center towards the tip of the root. The central root cap is clearly organized into several orderly horizontal tiers and longitudinal files of rectangular shaped cells (Dolan et al., 1993; Barlow, 2003; Wildwater et al.,

50 2005). The distal layers of root cap cells that get cast off periodically are constantly replenished by cell layers added proximally by cell division in the RAM (Dolan et al., 1993; Willemsen et al., 1998; Barlow, 2003). The central root cap cells that are placed vertically distal to the RAM are connected as continuous cell-tiers to the lateral root cap cells that extend laterally along the root tip (Dolan et al., 1993; Barlow, 2003).

2.2.4 Cell division in plant vegetative meristems and the role of sugars

In Arabidopsis, maintenance of stem cells in the SAM is mediated by a regulatory network that involves the negatively influencing ligand-receptor system CLAVATA (CLV) and the positively influencing homeodomain transcription factors WUSCHEL (WUS) and SHOOTMERISTEM-LESS (STM) (Laux et al., 1996; Long et al., 1996; Brand et al., 2000; Schoof et al., 2000). In the root, a similar mechanism to maintain stem cell fate has been proposed that involves the transcription factors SCARECROW (SCR), SHORTROOT (SHR) and WUSCHEL-RELATED HOMEOBOX-5 (WOX5) as the key players. These transcription factors maintain the initial cells of the root meristem in a division state and prevent their differentiation as proposed for WUS in the shoot (Sabatini et al., 1999; Kaya et al., 2001; Nawy and Benfey, 2001; Sabatini et al., 2003; De Veylder et al., 2007; Sarkar et al., 2007). In addition, environmental stimuli and phytohormones such as auxin, cytokinin, gibberellic acid (GA) and abscicic acid (ABA) are also involved in providing cell division cues to meristem cells in both the shoot and root meristems (den Boer and Murray, 2000; Leyser, 2003; Inze, 2005; De Veylder et al., 2007; Kyozuka, 2007). Cell division in the primary meristematic regions is a provider of the pool of cells required for plant growth, however the effects of cell division on plant development remain enigmatic. By using loss of function and overexpression studies as well as mechanical destruction of the meristematic tissue, some evidence has been generated that connects cell division in the shoot and root meristems and apex development, organogenesis and plant architecture (Loiseau, 1959; Laux et al., 1996; Van Lijsebettens

51 and Clarke, 1998; Nawy and Benfey, 2001; de Jager et al., 2005; Xu et al., 2008; Wu et al., 2005; Skylar et al., 2011).

Depending upon the stimuli received, cells in the G1 phase in plant meristems could progress into S phase with the initiation of DNA synthesis (i.e. behave as a ‘stem cell’) or exit the cell cycle to adopt a mitotically inactive quiescent state (G0), differentiate or undergo programmed cell death (den Boer and Murray, 2000). Transcriptional induction of Cyclin D by mitogens including cytokinin, brassinosteroids and auxin is shown to promote transition of cells from G1 to S phase to cause commitment to cell division (Dewitte and Murray, 2003; Gutierrez et al., 2002;). Further, Cyclin D is considered a rate-limiting factor in G1/S transition (Menges et al., 2006). Events downstream of Cyclin D induction lead to phosphorylation of the cell division inhibitors retinoblastoma- related proteins, which upon phosphorylation dissociate from the E2F family of transcription factors thereby abolishing the inhibitory effect. The freed E2F transcription factors bind regulatory sequences in the E2F-resonsive S-phase genes, thereby committing to cell division (de Jager and Murray, 1999; den Boer and Murray, 2000; Riou-Khamlichi et al., 2000). S-phase genes that contain E2F binding sequences in Arabidopsis include genes coding for origin recognition complex (ORC) subunits of DNA replication (Diaz-Trivino et al., 2005), other components of the pre-replication complex that licenses DNA for replication including AtCDC6 (de Jager et al., 2001; Ramos et al., 2001), and a minichromosome maintenance gene (MCM3) (Stevens et al., 2002) as well as condensin genes AtCAP-E1 and AtCAP-E2 required for maintenance of chromosomal integrity during cell division (Siddiqui et al., 2003).

The ability of sucrose, the major transported sugar resulting from photosynthesis to induce plant mitosis has been shown since early times (Ballard and Wildman, 1964; Van't Hof, 1966; Van't Hof and Rost, 1972). Sucrose induces Cyclins D1, D2 and D3 transcription and likely involves a phosphorelay-type signaling mechanism independent of de novo protein synthesis (Cockcroft et al., 2000; Gaudin et al., 2000; Riou-Khamlichi

52 et al., 2000; Dewitte and Murray, 2003). In cell cultures, Cyclin D induction by sucrose has been shown to occur via a cytokinin-independent signal transduction pathway (Riou- Khamlichi et al., 2000). However, in intact plants evidence of cross-talk between sucrose signaling and phytohormones including cytokinin is emerging (Eveland and Jackson, 2011). By sucrose depletion in the medium of suspension-cultured Arabidopsis cells as well as cultured primary root tips of sunflower (Helianthus annuus) and pea (Pisum sativum), it has been shown that the majority of the cells arrest in the G1 phase in response to sucrose withdrawal and that re-supply of sucrose leads to progression to the S-phase via resumption of cell division (Van't Hof, 1966; Van't Hof and Rost, 1972; Riou-Khamlichi et al., 2000). Thus, sucrose has been associated with G1/S transition in the cell cycle via Cyclin D induction. Similar effects as sucrose have been shown in response to glucose supplementation as well (Riou-Khamlichi et al., 2000). However, Arabidopsis mutants defective in G1/S transition upstream of Cyclin D induction have not been identified yet. Stress signaling mediated by ABA has been shown to negatively regulate Cyclin D activity via a poorly understood mechanism (den Boer and Murray, 2000). Thus, despite the central role played by Cyclin D in regulating likely the main control point to cell cycle entry (i.e. G1/S transition), the signaling pathways that connect environmental cues, hormones and nutrients to the regulation of Cyclin D levels, interconnections between these cascades and the inhibitory mechanisms involved remain elusive in plants (den Boer and Murray, 2000; Gaudin et al., 2000; Riou-Khamlichi et al., 2000; Gutierrez et al., 2002; Richard et al., 2002).

Novel findings regarding G1/S-independent sucrose-mediated rescue were reported by Skylar et al., 2011 and Wu et al., 2007 during further studies on the mutant of the homeobox transcription factor, STIMPY (STIP) (Wu et al., 2005). The stip mutant showed vegetative phenotypes of defective shoot and root apex development which were rescued by sucrose supplementation (Wu et al., 2005). This suggested possible G1/S arrest in the cell cycle for stip mutation (Wu et al., 2005). However, further studies showed that cell cycle arrest in stip mutant occurred in the G2 phase and was rescued by

53 sucrose supplementation. According to Skylar et al., 2011, the stip mutation induces expression of a repressor (i.e. TPR-DOMAIN SUPPRESSOR OF STIMPTY; TSS) of a G2-phase associated positive regulator of the cell cycle while sucrose supplementation downregulates this repressor in a stip mutation-dependant manner to suppress the stip mutation.

2.2.5 Common phenotypes of cell cycle mutants and wild-type plants that were induced for cell division arrest

The characterized Arabidopsis mutants defective in cell cycle progression are limited but include mutants carrying mutations in genes involved in different phases of the cell cycle including S and G2 functions; STIMPY (STIP) (Wu et al., 2005), FASCIATA (FAS) (Kaya et al., 2001), Nucleosome Assembly Protein1-Related Protein (NRP) (Zhu et al., 2006), TEBICHI (TEB) (Inagaki et al., 2006), Meiotic Recombination 1 (MRE1) (Bundock and Hooykaas, 2002), AtCDC5 (Lin et al., 2007), RPA2A/ROR1 (Xia et al., 2006), BRUSHY1 (BRU1) (Takeda et al., 2004), INCURVATA2 (ICU2) (Barrero et al., 2007), Ribonucleotide reductase (RNR) (Wang and Liu, 2006), M phase functions; HOBBIT (HBT) (Willemsen et al., 1998; Blilou et al., 2002), Chromatin Associated Protein Subunit E (AtCAP-E1 and AtCAP-E2) (Siddiqui et al., 2003), CCS52A2 (Van Straelen et al., 2009) and cytokinesis functions; TPLATE (Van Damme et al., 2006). In addition, deficiency in γ-Tubulin proteins (TUBG1 and TUBG2) that are important for the assembly of spindle, phragmoplast and cortical microtubule arrays also show cell division defects (Binarova et al., 2006; Pastuglia et al., 2006). The descriptions of phenotypes of these mutants are largely limited to the particular questions addressed and the organs investigated in a given study and therefore do not provide explicit details on the temporal and spatial development of all phenotypes of the mutant plants. Nevertheless, commonality in certain phenotypes can be discerned among these mutants irrespective of their phase or mechanism of arrest in the cell cycle. These common phenotypes are valuable tools in understanding the broader impact of cell division on

54 plant development and for dissecting the signaling and molecular mechanisms underlying the manifested phenotypes.

Stochastic nature of phenotypes leading to high variability in the severity of defects in different seedlings of the same population is commonplace for cell cycle mutants (Kaya et al., 2001; Bundock and Hooykaas, 2002; Takeda et al., 2004; Wu et al., 2005). While retarded seedling growth is commonly reported, lethality of the most severely affected seedlings is also observed in some of these mutants (Willemsen et al., 1998; Bundock and Hooykaas, 2002; Wu et al., 2005; Binarova et al., 2006; Pastuglia et al., 2006; Lin et al., 2007; Vanstraelen et al., 2009; Van Damme et al., 2006). The most common shoot phenotypes include delayed and/or distorted primary/true leaves, halt of initiation of true leaves after the development of a few (Kaya et al., 2001; Blilou et al., 2002; Bundock and Hooykaas, 2002; Siddiqui et al., 2003; Binarova et al., 2006; Inagaki et al., 2006; Pastuglia et al., 2006; Wang and Liu, 2006; Barrero et al., 2007; Lin et al., 2007) and the arise of multiple shoots from the shoot apex following failure of the development of the primary meristem (Bundock and Hooykaas, 2002; Takeda et al., 2004; Binarova et al., 2006; Wang and Liu, 2006; Barrero et al., 2007; Lin et al., 2007; Vanstraelen et al., 2009). At the cellular level, the cell organization in the SAM and true leaves appear disrupted with aberrant cell divisions and commonly lacking the organized cell files seen in wild- type plants (Kaya et al., 2001; Blilou et al., 2002; Siddiqui et al., 2003; Inagaki et al., 2006; Vanstraelen et al., 2009).

In the root, some common phenotypes include retarded root growth with differentiation of root hairs close to the root tip (i.e. thereby placing the root hair-zone adjacent to the cell division zone) and more prominent root hair development (Kaya et al., 2001; Blilou et al., 2002; Siddiqui et al., 2003; Takeda et al., 2004; Binarova et al., 2006; Inagaki et al., 2006; Pastuglia et al., 2006; Xia et al., 2006; Zhu et al., 2006; Lin et al., 2007; Vanstraelen et al., 2009; Van Damme et al., 2006). Additionally, disorganized cell files (Willemsen et al., 1998; Kaya et al., 2001; Bundock and Hooykaas, 2002; Siddiqui et al.,

55 2003; Inagaki et al., 2006; Zhu et al., 2006; Vanstraelen et al., 2009) and enlarged cells predominantly in the differentiation zone are visible in some of the reported cell cycle mutants (Siddiqui et al., 2003; Binarova et al., 2006; Pastuglia et al., 2006; Zhu et al., 2006) as well as in the inducible RanGAP RNAi lines (Xu et al., 2008). Wild-type plants that were induced chemically or by radiation for cell cycle arrest in G1, S and G2 phases also showed similar root phenotypes, including swollen cells in the differentiation zone (Culligan et al., 2004). The adult plant phenotypes of many of these mutants have shown to be variable and less documented, however, stunted and bushy phenotypes have been commonly reported (Bundock and Hooykaas, 2002; Takeda et al., 2004; Binarova et al., 2006; Wang and Liu, 2006; Xia et al., 2006; Vanstraelen et al., 2009).

In the literature, several of these phenotypes have apparently been considered hallmarks of mutants perturbed in S-phase functions including defective epigenetic gene silencing, DNA repair and replication and have aided in the identification of more S-phase defective mutants (Inagaki et al., 2006; Wang and Liu, 2006). These S-phase defective mutants constitute the largest representation among identified cell cycle mutants so far. However, these phenotypes resemble the phenotypes of the few characterized mutants carrying disrupted genes that function in other phases of cell cycle (Willemsen et al., 1998; Blilou et al., 2002; Wu et al., 2005; Binarova et al., 2006; Pastuglia et al., 2006; Wang and Liu, 2006) and with the wild-type plants that were treated to halt in G1 and G2 phases of the cell cycle as described above (Culligan et al., 2004). Therefore, these phenotypes appear to extend beyond arrests in the S-phase alone and seem common to cell cycle defects irrespective of the stage and mechanism of arrest. Taking this together with the evidence that connects proper cell cycle functioning with meristem and plant development (Laux et al., 1996; Van Lijsebettens and Clarke, 1998; Nawy and Benfey, 2001; de Jager et al., 2005; Wu et al., 2005; Skylar et al., 2011), I propose that at least some of these phenotypes are likely common downstream effects of aberrant cell division in the main meristems, irrespective of the phase or mechanism of arrest. The necessity for passage through certain phases of cell cycle for expression of cell differentiation factors

56 to cause proper differentiation has been suggested as a possibility in Arabidopsis development as seen in Drosophila neurons (Blilou et al., 2002). This could mean that perturbed cell division at any phase of the cell cycle could commonly affect certain developmental processes in a similar manner.

In this study, I have developed and analyzed T-DNA knockdown mutants of RanGAP in Arabidopsis and shown the requirement of RanGAP for cell division, proper meristem function and vegetative plant development using a severely RanGAP-deficient rg1-1/rg1- 1;rg2-2/rg2-3 mutant. The observed commonality of phenotypes between the RanGAP- deficient rg1-1/rg1-1;rg2-2/rg2-3 mutant, other cell cycle mutants and wild-type plants induced chemically or by radiation to perturb the cell cycle support a notion of common phenotypic hallmarks for aberrant cell cycle function irrespective of phase and mechanism of cell cycle arrest. This phenotypic index could potentially facilitate the identification of novel cell cycle-mutants and improve our understanding of the molecular mechanism of the cell cycle as well as effects of cell division on plant development.

2.3 Materials and Methods

2.3.1 Plant material, growth conditions and constructs

T3 or T4 bulk seeds of T-DNA insertion mutants rg1-1 (SALK_058630) and rg2-2 (SALK_006398) in Columbia ecotype (Col) were acquired from the Arabidopsis Biological Resource Center (ABRC). T-DNA mutant rg2-3 (FLAG_184A06) in Wassilewskija background (Ws) was acquired from the Versailles T-DNA lines collection (Bechtold et al., 1993; Bouchez et al., 1993). The position of the T-DNA insert was confirmed by sequencing to reside ~720 bp downstream of the start codon of RanGAP1 in rg1-1. In rg2-2 and rg2-3, the insert was confirmed to reside ~370 bp and

57 ~190 bp upstream of the start codon of RanGAP2, respectively. F1 populations containing RG1/rg1-1;RG2/rg2-2 and RG1/rg1-1;RG2/rg2-3 allelic combinations were obtained from crosses rg1-1/rg1-1;RG2/RG2 X RG1/RG1;rg2-2/rg2-2 and rg1-1/rg1- 1;RG2/RG2 X RG1/RG1;rg2-3/rg2-3 respectively. Subsequent self-fertilization produced F2 and F3 generations. Work leading to the production of F2 generation was carried out by Xu, X.M. rg1-1/rg1-1;rg2-2/rg2-2 was crossed as the female parent with pollen from rg1-1/ rg1-1;RG2/rg2-3 to obtain a trans-heterozygote rg1-1/rg1-1;rg2-2/rg2-3 and the co-segregating rg1-1/rg1-1;RG2/rg2-2 plants. Thus, both rg1-1/rg1-1;rg2-2/rg2-3 and rg1-1/ rg1-1;RG2/rg2-3 are Col/Ws hybrids.

Arabidopsis seedlings were grown under constant light on Murashige and Skoog (MS) plates [Per L: 4.3 g MS basal salt with macro- and micronutrients (Caisson Laboratories Inc., Rexburg, ID Cat # MSP0501), 1 ml of 1000X Gamberg B5 vitamin solution (Sigma Cat # G1019), 0.5 g MES, 8 g agar (USB Cat # 10906), pH 5.7] supplemented with the specified percentage of sucrose (0%, 1.5%, 2% or 3%) of weight (g) of sucrose per unit volume (ml) of medium [i.e. w/v]. In soil, the Arabidopsis plants were grown under standard long-day condition (16 h light and 8 h dark) with light intensity ~90-120 µmolm-2s-1, relative humidity ~50% and temperature ~22ºC. Light intensity was measured using BQM Quantum Light Meter (Hydrofarm Gardening Products). Temperature and humidity were measured using Onset HOBO data logger. On MS media, plants were grown under continuous light ~70-100 µmolm-2s-1, relative humidity ~50- 55% and temperature ~20-22ºC.

To incorporate transgenes into rg1-1/rg1-1;rg2-2/rg2-3 and co-segregating rg1-1/rg1- 1;RG2/rg2-2 plants, first transgenic rg1-1/rg1-1;rg2-2/rg2-2 plants were obtained by transformation of rg1-1/rg1-1;rg2-2/rg2-2 plants with the construct via floral dipping (Clough and Bent, 1998). Subsequently, rg1-1/rg1-1;rg2-2/rg2-2 carrying the transgene was used as the female parent to cross with rg1-1/ rg1-1;RG2/rg2-3 pollen to obtain the

58 progeny of rg1-1/rg1-1;rg2-2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 plants containing the transgene.

To obtain rg1-1/rg1-1;rg2-2/rg2-2 plants carrying the mitotic marker CycB1;1::GUS, the binary vector pBI101.3 containing the CycB1;1::GUS insert (a kind gift from John Celenza, Boston University, Massachusetts. Construct details are described in Colon- Carmona et al., 1999) was mobilized into the Agrobacterium strain GV3101 by electroporation. Subsequently, Arabidopsis rg1-1/rg1-1;rg2-2/rg2-2 plants were

transformed by floral dipping (Clough and Bent, 1998) and the primary transformants (T1 generation) were selected by Kanamycin resistance (50 µg/ml). Transgenic rg1-1/rg1- 1;rg2-2/rg2-2 + CycB1;1::GUS was subsequently crossed with pollen from rg1-1/rg1- 1;RG2/rg2-3 to obtain rg1-1/rg1-1;rg2-2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 plants containing the CycB1;1::GUS construct.

A transgenic line (Col background) containing a nuclear import marker; 35S-GFP-N7 (nuclear localization protein N7 fused to an N-terminal GFP tag and driven by CaMV 35S promoter - ABRC CS84731, GenBank accession number CAA16704, described by Cutler et al., 2000, Cutler and Somerville, 2005) was acquired from the ABRC and was crossed with rg1-1/rg1-1;rg2-2/rg2-2. F2 progeny was screened by PCR genotyping and GFP fluorescence to obtain transgenic rg1-1/rg1-1;rg2-2/rg2-2 + 35S-GFP-N7, which was subsequently crossed with pollen from rg1-1/rg1-1;RG2/rg2-3 to obtain rg1-1/rg1- 1;rg2-2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 plants containing 35S-GFP-N7.

To obtain Arabidopsis mutants with a perturbed Ran cycle, inducible GDP-locked or GTP-locked Ran1 constructs were developed. For this purpose, dominant negative point mutations were introduced to Arabidopsis Ran1 with the QuickChange Site-Directed Mutagenesis kit (Stratagene, La Jolla, CA). Thr27 in Arabidopsis Ran1 (corresponding to Thr24 in human Ran) was mutated to Asn to construct a nucleotide-free/GDP-locked form designated Ran1T27N similar to human RanT24N (Klebe et al., 1995). Similarly,

59 Gln72 of Arabidopsis Ran1 (corresponding to Gln69 in human Ran) was mutated to Leu to obtain the Ran1Q72L mutant which resembles the human RanQ69L mutant that fails to hydrolyze GTP and thereby adopts a GTP-locked conformation (Bischoff et al., 1994). HA-Ran1 (control), HA-Ran1T27N and HA-Ran1Q72L were then recombined into the destination vector pMDC7 for estrogen-inducible expression in plants (Curtis and Grossniklaus, 2003). Transgenic Arabidopsis plants were selected on MS plates containing Hygromycin (35 µg/ml) and Cefataxin (125 µg/ml) after Agrobacterium- mediated transformation by floral dip method (Clough and Bent, 1998). For induction, seedlings were either germinated directly on or transferred to MS plates containing β- estradiol (10 µM, Sigma, for inducible expression), while ethanol (solvent) was used for control (i.e. uninduced) treatments. Development of Arabidopsis Ran mutants and experiments thereof were conducted by Xu, X.M.

2.3.2 PCR-based genotyping of T-DNA insertion lines

F2 and F3 generations were screened by PCR genotyping (http://signal.salk.edu/tdnaprimers.2.html) to identify mutant and wild-type alleles of RanGAP. Genomic DNA was extracted as described (Krysan et al., 1999). Details of the primers used are summarized in Table 1.1. For screening of the rg2-2 allele 032721FP and LBa1 primers were used.

2.3.3 Immunoblot analysis

For immunoblot analysis to compare RanGAP2 expression level between wild-type (Col) and rg1-1/rg1-1;rg2-2/rg2-2 (Col), tissues were harvested from seedlings of 8 days post- germination (8 dpg). For the comparative immunoblot to analyze RanGAP2 expression level between rg1-1/rg1-1;rg2-2/rg2-2 and rg1-1/rg1-1;rg2-2/rg2-3, 12 dpg rg1-1/rg1- 1;rg2-2/rg2-2 seedlings containing a hybrid background of approximately three quarters of Col and one quarter of Ws background (i.e. Col:Ws ~3:1) similar to rg1-1/rg1-1;rg2-

60 2/rg2-3 were obtained. For this, the rg1-1/rg1-1;RG2/rg2-2 plants (Col: Ws ~3:1) that were obtained along with rg1-1/rg1-1;rg2-2/rg2-3 (Col: Ws ~3:1) [i.e. by the cross ♀rg1-1/rg1-1;rg2-2/ rg2-2 (Col) X ♂rg1-1/ rg1-1;RG2/rg2-3 (Col: Ws 1:1)] were selfed and the progeny was genotyped to identify rg1-1/rg1-1;rg2-2/rg2-2 plants which would contain a hybrid background of Col: Ws ~3:1. Seedlings of rg1-1/rg1-1;rg2-2/rg2- 3 and rg1-1/rg1-1;rg2-2/rg2-2 with the hybrid background Col: Ws ~3:1 were grown for 12 days in continuous light on MS plates and a pool of whole seedlings for each genotype was used for the immunoblot analysis. Anti-RanGAP1 and Anti-RanGAP2 antibodies were described in (Jeong et al., 2005) and Chapter 1, respectively. Anti-MFP1 antibody (OSU91) described in (Jeong et al., 2003) was used to detect the expression of endogenous MAR binding filament-like protein 1 (MFP1) as the loading control.

2.3.4 Embryo observations

Mature green seeds of siliques from floral development stage 17 (see Table 1.2 for details on floral development stages) were mounted on clearing solution (chloral hydrate 8 g: glycerol 1 ml: water 2 ml) and pressure was applied using the cover slip to release embryos from the seeds. Embryos were allowed to clear for more than 6 hrs at room temperature and observed with DIC microscopy.

2.3.5 Scanning electron microscopy

Tissues were fixed in 3% Glutaraldehyde in 0.1 M potassium phosphate buffer (pH 7.2) at room temperature for 4 h and subsequently at 4°C for 48 h. After washing with 0.1 M potassium phosphate buffer (pH 7.2), they were dehydrated 20 min each in a series of ethanol concentrations (v/v) of 25% 50%, 70%, 80%, 85%, 90%, 95% and 3 times in 100%. Subsequently they were treated for 20 min each with 25%, 50% and 75% Hexamethyldisilazane (HMDS) in ethanol followed by 3 times in 100% HMDS. Treated samples were mounted on metal stubs, air dried overnight, sputter coated with gold and

61 observed in FEI Nova 400 NanoSEM (Campus Microscopy and Imaging Facility, The Ohio State University)

2.3.6 Sectioning

Tissues suspended in 2.8% (v/v) glutaraldehyde in 0.1 M HEPES buffer (pH7.2) and 0.02% (v/v) Triton X-100 were vacuum infiltrated at 15 InchHg for 30 min and incubated at room temperature for 4 h and subsequently at 4°C for 48 h. After washing with a buffer containing 0.1 M HEPES (pH7.2) and 0.02% (v/v) Triton X-100, the tissues were

post-fixed in 1% OsO4 in 0.1 M HEPES buffer (pH7.2) and 0.02% (v/v) Triton X-100 at 4°C for 24 h in a dark container. After buffer wash, tissues were dehydrated 20 min each through an ascending acetone series of 20%, 30%, 40%, 50% 60%, 70%, 80%, 90% and 3 times in 100% and embedded in the Low Viscosity Embedding Media Spurr’s Kit

(Electron Microscopy Sciences, Hatfield, PA) according to the manufacturer’s protocol. 0.5 µm-2 µm sections were obtained using the ultramicrotome (Campus Microscopy and Imaging Facility, The Ohio State University) and stained with 1% aqueous toluidine blue containing 0.5% pyronin.

2.3.7 Complementation assay

The native promoter-RanGAP1 and native promoter-RanGAP2 constructs used for the complementation assay were generated and mobilized into the Agrobacterium strain GV3101 as described in Chapter 1. Subsequently, Arabidopsis rg1-1/rg1-1;rg2-2/rg2-2 plants were transformed with each construct by floral dipping (Clough and Bent, 1998)

and the primary transformants (T1 generation) were selected by hygromycin resistance (30 µg/ml). Siliques of T1 transformants were evaluated for complementation. Transgenic T1 rg1-1/rg1-1;rg2-2/rg2-2 containing the native promoter construct was subsequently crossed with pollen from rg1-1/rg1-1;RG2/rg2-3 to obtain rg1-1/rg1-1;rg2- 2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 plants containing each native promoter-driven

62 construct. Three T1 lines of transgenic rg1-1/rg1-1;rg2-2/rg2-2 plants carrying each native promoter construct were used for the crosses. Phenotypic observations of seedlings were conducted on 2% sucrose (w/v) MS plates under continuous light to assay for complementation and the seedlings were genotyped using PCR for confirmation. An immunoblot analysis was performed to assay the expression of the transgene in phenotypically recovered seedlings compared to non-transformed rg1-1/rg1-1;rg2-2/rg2- 3 and wild-type.

2.3.8 β-Glucuronidase assay

Histochemical staining for β-Glucuronidase (GUS) activity in seedlings was performed as described in Chapter 1. In order to improve the clarity of the stained shoot apical meristem specimens, seedling shoots were cleared with a clearing solution (chloral hydrate 8 g: glycerol 1 ml: water 2 ml) after GUS staining as described in Chapter 1.

2.3.9 Sucrose assay

To study the effect of sucrose on the mutant growth and development, rg1-1/rg1-1;rg2- 2/rg2-3 and the co-segregating rg1-1/rg1-1;RG2/rg2-3 plants were grown in MS plates containing 0%, 1.5%, 2% and 3% sucrose (w/v) under constant light for ~2 weeks to assay seedling phenotypes. Some seedlings were subsequently transferred to soil or a hydroponic set-up containing a liquid MS medium of the same composition to assay adult plant phenotypes. The identity of rg1-1/rg1-1;rg2-2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 plants resulting from the segregating pool of seeds and used in the sucrose assay was confirmed by PCR-based genotyping.

63 2.3.10 Confocal microscopy

For staining of cell boundries in root tips, propidium iodide and FM4-64 stains were used in water at final concentrations of 25 µg/ml and 5 µM respectively. Confocal microscopy images of propedium iodide/FM4-64 staining and GFP-N7 localization in root tips were collected on a PCM 2000/Nikon Eclipse E600 confocal laser scanning microscope as described in Rose and Meier, 2001.

2.4 Results

2.4.1 RanGAP knockdown rg1-1/rg1-1;rg2-2/rg2-2 plants showed mild defects in vegetative growth and development

The cross between RanGAP1 knockout single mutant; rg1-1/rg1-1;RG2/RG2 and RanGAP2 knockdown single mutant; RG1/RG1;rg2-2/rg2-2 yielded viable rg1-1/rg1- 1;rg2-2/rg2-2 plants in the F2 generation. The position of T-DNA insert in the rg1-1 allele and the immunoblot indicating rg1-1 as a null allele were shown in Figures 1.1A and 1.2A respectively. The position of the T-DNA insert in the rg2-2 allele is shown in Figure 2.1A. The rg2-2 allele was shown to be a RanGAP2 knockdown allele by immunoblot analysis (Figure 2.1B). Primers used for genotyping to identify rg1-1/rg1- 1;rg2-2/rg2-2 plants are described in 1.3.2 and 2.3.2.

Because the RanGAP knockout rg1-1/rg1-1;rg2-3/rg2-3 individuals are female gametophytic lethal (Rodrigo-Peiris et al., 2011), knockdown T-DNA double mutant; rg1-1/rg1-1;rg2-2/rg2-2 plants were phenotypically analyzed in soil and MS medium containing 1.5% and 2% sucrose (w/v) (growth conditions are described in Materials and methods). The rg1-1/rg1-1;rg2-2/rg2-2 mutant was very similar to wild-type both at the seedling and mature plant level (Figure 2.2A and B), with a possible mild delay in

64 seedling development (Figure 2.2A). A short silique phenotype was observed in the rg1- 1/rg1-1;rg2-2/rg2-2 mutant and the siliques contained a reduced number seeds (Figure 2.3A). Variable integument defects in ovules, and female gametophytic arrests at different stages of development were observed (Figure 2.3B-G). Native promoter-driven RanGAP constructs complemented these phenotypes (RG1 complementation of reproductive phenotypes is shown in Figure 2.3H and I).

2.4.2 Severe RanGAP knockdown trans-heterozygous rg1-1/rg1-1;rg2-2/rg2-3 mutant seedlings showed prominent shoot and root phenotypes

Progeny of the genetic cross between ♀rg1-1/rg1-1;rg2-2/rg2-2 X ♂rg1-1/rg1- 1;RG2/rg2-3 was analyzed for phenotypes. Mature embryos resulting from the cross were uniform when observed by DIC light microscopy (a representative embryo is shown in Figure 2.4A) suggesting that rg1-1/rg1-1;rg2-2/rg2-3 embryos in the progeny are indistinguishable from the co-segregating rg1-1/rg1-1;RG2/rg2-2 embryos. When seeds resulting from the ♀rg1-1/rg1-1;rg2-2/rg2-2 X ♂rg1-1/rg1-1;RG2/rg2-3 were germinated in soil and MS medium containing a 1.5% and 2% sucrose (w/v) (growth conditions are described in Materials and methods), defects in seed germination efficieny was not observed among the progeny suggesting that rg1-1/rg1-1;rg2-2/rg2-3 germinated with a similar efficiency to rg1-1/rg1-1;RG2/rg2-2. An observation not further quantitatively analysed was that ~15% of these seeds germinated with the shoot emerging prior to the root radicle which were identified as rg1-1/rg1-1;rg2-2/rg2-3 by genotyping and/or seedling phenotypes as described below.

The seedlings of the progeny of ♀rg1-1/rg1-1;rg2-2/rg2-2 X ♂rg1-1/rg1-1;RG2/rg2-3 showed an approximate 1:1 ratio between seedlings of wild-type-like morphology and a mutant with growth and development phenotypes (mutant phenotypes are described below) when grown in soil and MS medium containing a 1.5% and 2% sucrose (w/v) [wild-type-like, 26; mutant phenotype, 30; χ²(1, N = 56) = 0.286; 0.7 >p >0.5 for the

65 probability that a deviation from a 1:1 ratio is due to chance; N refers to the number of individuals analyzed]. Genotyping for RG2 and rg2-3 alleles showed that the mutant phenotype was associated with rg1-1/rg1-1;rg2-2/rg2-3 and that the wild-type-like plants were rg1-1/rg1-1;RG2/rg2-2 (Figure 2.4B). Thus the clear split between large, wild-type- like seedlings (rg1-1/rg1-1;RG2/rg2-2) and small, underdeveloped seedlings (rg1-1/rg1- 1;rg2-2/rg2-3) enabled non-destructive discrimination between the two genotypes for the subsequent analyses. Immunoblot analysis suggested that rg1-1/rg1-1;rg2-2/rg2-3 mutants contained a further knockdown level of RanGAP compared to rg1-1/rg1-1;rg2- 2/rg2-2 plants (Figure 2.4C).

Phenotypic differences between rg1-1/rg1-1;RG2/rg2-2 and rg1-1/rg1-1;rg2-2/rg2-3 were analyzed in MS medium containing 2% sucrose (w/v) [and were recapitulated in MS medium containing 1.5% sucrose (w/v) and in soil]. In comparison to rg1-1/rg1- 1;RG2/rg2-2 plants, rg1-1/rg1-1;rg2-2/rg2-3 showed post-germination seedling growth and developmental phenotypes with a high degree of variation in the phenotypic severity between individual seedlings, and in the morphology of shoot apex phenotypes (described below). Growth retardation in rg1-1/rg1-1;rg2-2/rg2-3 seedlings was observable from ~2 dpg resulting in small shoots, short roots and overall smaller seedling sizes (Figure 2.5A and B). In the shoot apex of rg1-1/rg1-1;rg2-2/rg2-3, reduced expansion of the cotyledons was observed (Figure 2.5B). Development of the shoot apex was perturbed (Figure 2.6) including a ~2-5 dpg delay in the initiation of the first pair of true leaves. The morphology of first pair of true leaves was generally atypical where they ceased to grow after initiation (Figure 2.7A) or often developed crinkled (Figure 2.7B), oblong or bifid leaves (Figure 2.7C) or sometimes developed into rod-like structures that lack dorso-ventral cell specialization (Figure 2.7D). Note the phenotypic variation between individual rg1-1/rg1-1;rg2-2/rg2-3 seedlings in Figure 2.7.

The cellular organization of the shoot apex was disturbed in rg1-1/rg1-1;rg2-2/rg2-3 seedlings compared to rg1-1/rg1-1;RG2/rg2-2 (Figure 2.8). Particularly the layered

66 organization of the cell files observed in the first true leaves of rg1-1/rg1-1;RG2/rg2-2 seedlings was disturbed in rg1-1/rg1-1;rg2-2/rg2-3. Irregular cell shapes, sizes and cell division planes were observable in the first true leaves of rg1-1/rg1-1;rg2-2/rg2-3. The cellular organization of the shoot apex was disrupted by the random occurrence of protuberances containing cell masses, some of which were identifiable by gross morphology as containing meristem-like organization (i.e. cell layer arrangement resembling meristem structure) (Figure 2.8B).

A period of severely retarded shoot apex development occurred in rg1-1/rg1-1;rg2-2/rg2- 3 seedlings noticeable from ~2-3 dpg to ~8-12 dpg during which time the slow expansion of the cotyledons, malformation of the first true leaves and enlargement of the protuberances in the shoot apex took place. The most severely affected seedlings died (i.e. non-regenerating seedlings: ~25%) during this retarded growth phase with some of them producing terminal glandular or callus like structures at the shoot apex (Figure 2.9A and B). Of the surviving seedlings, some of the protuberances in the shoot apex were identifiable as adventitious shoot primordia at ~8-12 dpg which were varied in number between the individual seedlings (~2-5 shoot primordia/apex) with each primordium containing a meristem surrounded by leaf primordia (Figure 2.9C). In addition to the malformed first true leaves and adventitious shoot primordia in rg1-1/rg1-1;rg2-2/rg2-3, additional leaves continued to develop in isolated places of the shoot apex (Figure 2.9C). Thus, while rg1-1/rg1-1;RG2/rg2-2 showed an organized shoot apex with the true leaves arranged in an orderly radial pattern, the shoot apex of rg1-1/rg1-1;rg2-2/rg2-3 was highly disorganized (2.9C and 2.9D). Adventitious shoot meristems of rg1-1/rg1-1;rg2- 2/rg2-3 developed into small rosettes collectively forming a compact shoot (Figure 2.9E- G).

Morphological and anatomical alterations were also observed in the short seedling roots of rg1-1/rg1-1;rg2-2/rg2-3. Root hairs developed closer to the root tip in rg1-1/rg1- 1;rg2-2/rg2-3, were generally more abundant and longer compared to rg1-1/rg1-

67 1;RG2/rg2-2 (Figure 2.10A and B). Meristematic and elongation zones appeared shorter in rg1-1/rg1-1;rg2-2/rg2-3 roots compared to rg1-1/rg1-1;RG2/rg2-2 (Figures 2.10A and B, 2.16C and D). In the root cap region of rg1-1/rg1-1;rg2-2/rg2-3 irregular cell shapes, obliquely placed cell walls and disorganized cell files were observed (Figures 2.10C and E). Disturbed organization of cell files was observed in the non-root cap ground tissue (ground tissue defined as cortex and endodermis according to Paquette and Benfey, 2005) of some rg1-1/rg1-1;rg2-2/rg2-3 roots. Cell file disorganization in the ground tissue was highly variable between seedlings and depending on seedling age (one example is shown in Figure 2.10C).

2.4.3 rg1-1/rg1-1;rg2-2/rg2-3 showed adult vegetative and reproductive phenotypes

Short reproductive shoots were initiated from the adventitious rosettes of rg1-1/rg1- 1;rg2-2/rg2-3 when grown in soil from the seed stage and in plants transferred to soil after ~2 weeks of growth on 1.5% or 2% sucrose media (Figure 2.11A-C). Reproductive shoots of rg1-1/rg1-1;rg2-2/rg2-3 showed normal phyllotaxy (Figure 2.11A-C). The floral morphology was unaltered, but a reduced number of flowers per inflorescence was observed (i.e ~4-5 flowers per inflorescence in rg1-1/rg1-1;rg2-2/rg2-3 compared to ~7- 8 in rg1-1/rg1-1;RG2/rg2-2). Mature plants were short and bushy with a compact basal shoot consisting of the multiple rosettes (Figure 2.11D). The size of siliques was significantly smaller in rg1-1/rg1-1;rg2-2/rg2-3 consisting predominantly of aborted female gametophytes and rarely containing fertile seeds (~0-10 fertile seeds per plant) (Figure 2.11E, Table 2.1). The number of ovule primordia initiated per silique was reduced by ~50% in rg1-1/rg1-1;rg2-2/rg2-3 compared to rg1-1/rg1-1;RG2/rg2-2 (Table 2.1). The reproductive shoots of rg1-1/rg1-1;rg2-2/rg2-3 branched excessively and continued to flower and fruit beyond the life span of rg1-1/rg1-1;RG2/rg2-2 plants that have terminated flowering/fruiting and were in the path of senescence (Figure 2.11D). A mild phenotype in the performance of rg1-1;rg2-3 pollen was observed in comparison to

68 rg1-1;rg2-2 pollen developed on rg1-1/rg1-1;rg2-2/rg2-3 mother plants. When wild-type (Col) pistils were pollinated with pollen from rg1-1/rg1-1;rg2-2/rg2-3, the knockout rg2- 3 allele showed only 77.5% genetic transmission through the pollen compared to the 100% genetic transmission rate of the competing knockdown rg2-2 allele (Table 2.2) (allelic/genetic transmission as described in Howden et al., 1998).

2.4.4 RanGAPs show high expression in the shoot and root meristems

To assay temporal and spatial expression pattern of RanGAP1 and RanGAP2 in vegetative plant, publicly available microarray data (GENEVESTIGATOR; http://www.genevestigator.ethz.ch; Zimmermann et al., 2004) were analyzed. Expression (log2>10) for both RanGAPs was reported for all developmental stages from germination to post-fruiting. Figure 2.12 illustrates a higher level of RanGAP1 and RanGAP2 expression in the shoot apex and primary root meristem in the vegetative plant. Note that RanGAP1 and RanGAP2 are expressed (log2 >9.0) in all anatomical locations analyzed.

To further analyze RanGAP expression with respect to anatomical locations, transgenic RanGAP1::GUS and RanGAP2::GUS seedlings were assayed. A high level of GUS expression was detected in the primary SAM, primary RAM, lateral root initiating cells (i.e. meristematic cells in the pericycle) and in lateral RAMs in transgenic lines carrying RanGAP1::GUS and RanGAP2::GUS constructs (Figure 2.13). Note that RanGAP expression is not limited to vegetative meristem alone. (RanGAP::GUS data were generated by Wang, H-J.).

2.4.5 Shoot and root phenotypes were complemented with RanGAP1 and RanGAP2 genomic constructs

To evaluate whether the observed phenotypes were caused by RanGAP deficiency, functional complementation with RanGAP1 and RanGAP2 genomic constructs was

69 performed. Seedling phenotype was rescued by complementation with RanGAP1 and RanGAP2 genomic constructs independently (Figure 2.14, RanGAP2 data not shown). This confirms the cause of these phenotypes as a deficiency in RanGAP and that the two RanGAP paralogs are redundant for the underlying function(s). Details of constructs and complementation assay are as described in Materials and methods.

2.4.6 Phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 caused by depletion of RanGAP is likely independent from Ran cycle-based nuclear transport

To analyze whether the phenotypes observed in rg1-1/rg1-1;rg2-2/rg2-3 resulted from a downstream effect of disrupted nuclear transport due to RanGAP deficiency, the efficiency of nuclear import of a nuclear import marker was assayed as a preliminary study. Here, the subcellular localization of the GFP-fused nuclear imported protein N7 (GFP-N7, Cutler et al., 2000; Cutler and Somerville, 2005) was evaluated in transgenic rg1-1/rg1-1;rg2-2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 plants grown on 2% sucrose (w/v) MS medium. A defect in GFP-N7 nuclear import was not observed for rg1-1/rg1-1;rg2- 2/rg2-3 in comparison to rg1-1/rg1-1;RG2/rg2-2 plants (Figure 2.15A-D). Figure 2.15E shows that GFP-N7 import in the inducible expression lines of a GDP-locked Ran mutant (Ran1T27N) and a GTP-locked Ran mutant (Ran1Q72L) (described in Materials and methods) is impeded, which provides a positive control to comparatively evaluate the efficiency of GFP-N7 nuclear import in rg1-1/rg1-1;rg2-2/rg2-3 [Figure 2.15E was generated by Xu, X.M.]. A second nuclear import protein GFP-WIP1ΔTDF (Xu et al., 2007a) also showed impeded nuclear import in the inducible GDP-locked and GTP- locked mutant lines (data generated by Xu, X.M. and not shown here). In contrast, nuclear import defects were not observable in the root tip cells (including cells with irregularly placed cell walls and the severely affected swollen cells) of inducible RanGAP RNAi lines described in Xu et al., 2008 using GFP-WIP1ΔTDF marker (Figure 2.15F-I, data generated by Xu, X.M.) and GFP-N7 marker (data generated by Xu, X.M.

70 and not shown here). Results for only the RanGAP2RNAi/rg1-1 RanGAP RNAi line is shown in Figure 2.15F-I, which were recapitulated for RanGAP1RNAi/rg2-3 line as well.

2.4.7 Cell division is restricted to compact regions in the shoot and root apices of rg1-1/rg1-1;rg2-2/rg2-3

The seedling and adult phenotypes observed in rg1-1/rg1-1;rg2-2/rg2-3 showed similarity to documented cell-cycle mutants irrespective of the phase or mechanism of the arrest in cell cycle, as well as to wild-type seedlings that were induced chemically or by radiation to arrest in G1, S or G2 phases of the cell cycle (see Introduction for details of these phenotypes). Therefore, to assay mitotic activity of the shoot and root apices of rg1- 1/rg1-1;rg2-2/rg2-3, transgenic plants containing the mitotic marker CycB1;1:GUS (Colon-Carmona et al., 1999) were evaluated. CycB1;1::GUS expression indicates regions with cell division activity.

Expression of CycB1;1:GUS evaluated by β-Glucuronidase (GUS) assay was restricted to compact regions in the shoot apex of non-regenerating rg1-1/rg1-1;rg2-2/rg2-3 + CycB1;1:GUS seedlings compared to rg1-1/rg1-1;RG2/rg2-3 + CycB1;1:GUS seedlings (Figure 2.16A and B). Thus, the reduced area of cell division concurred with the reduced area in the SAM and small shoot apex size in rg1-1/rg1-1;rg2-2/rg2-3 seedlings. However, the shoot apex of seedlings in the process of regeneration by producing adventitious shoot primordia showed intense staining in a larger area of shoot apex likely due to the meristematic activity of the multiple regenerating adventitious shoot primordia in the shoot apex (data not shown). Consistent with the reduced length of root meristematic and elongation zones in rg1-1/rg1-1;rg2-2/rg2-3 seedlings, CycB1;1:GUS expression was restricted to a compact region towards the apex in rg1-1/rg1-1;rg2-2/rg2- 3 + CycB1;1:GUS primary root compared to rg1-1/rg1-1;RG2/rg2-3 + CycB1;1:GUS (Figure 2.16C and D). Because the elongation zone of the root contains a portion of dividing cells in addition to the meristematic zone (Schiefelbein and Benfey, 1994;

71 Siddiqui et al., 2003), CycB1;1:GUS expression is observed in both meristematic and elongation zones.

2.4.8 Sucrose supplementation complements phenotypes of rg1-1/rg1-1;rg2-2/rg2- 3 seedlings

Based on 1) the findings in cell culture of Arabidopsis, and root tip cultures of sunflower and pea that sucrose supplementation could rescue G1-arrested cells (see Introduction), and 2) the cell cycle-deficient phenotypes observed in RanGAP mutants, I set out to test whether sucrose supplementation could rescue cell division and cause phenotypic recovery in rg1-1/rg1-1;rg2-2/rg2-3 plants. For this purpose, the progeny of ♀ rg1-1/rg1- 1;rg2-2/rg2-2 X ♂ rg1-1/rg1-1;RG2/rg2-3 segregating for rg1-1/rg1-1;rg2-2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 were germinated on MS medium containing 0% or 3% sucrose (w/v) and analyzed for phenotypes.

In contrast to ~100% of the seedlings that showed wild-type-like development up to ~5 dpg on 3% sucrose medium, only ~50% of the seedlings were wild-type-like on 0% sucrose medium. Among the other ~50% of the seedlings on 0% sucrose medium that showed retarded development, the majority (~90%) halted growth at the stage of two cotyledons prior to seedling establishment (Figure 2.17A) (‘seedling establishment’ defined as the formation of true leaves and root elongation according to Cernac et al., 2006). These seedlings did not undergo further development and underwent seedling death. The remainder of the retarded seedlings on 0% sucrose medium (i.e. ~10%) was able to escape the two-cotyledon stage and develop into severely retarded seedlings that underwent death within a few days after seedling establishment. Genotyping of representative seedlings associated the two-cotyledonary phenotype with rg1-1/rg1- 1;rg2-2/rg2-3 and the wild-type-like phenotype with rg1-1/rg1-1;RG2/rg2-2 (Figure 2.17A - bottom panel).

72 On 3% sucrose medium, ~50% of the seedlings showed delayed development of the first true leaf pair by about 1-2 days at ~5 dpg when observed by naked eye leading to an apparent temporary halt of shoot growth and subsequent recovery (i.e. ‘stop-and-go’ shoot development phenotype: Laux et al., 1996). The recovered shoots resembled wild- type-like shoots in phyllotaxy but showed a marginal retardation compared to the co- segregating wild-type-like plants when observed at 12 dpg (Figure 2.17B). Similarly, root growth rate and length showed a mild retardation. Genotyping of representative seedlings showed that the ‘stop and go’ and mildly retarted growth phenotype was associated with rg1-1/rg1-1;rg2-2/rg2-3 (Figure 2.17B - bottom panel). The delayed development of the first true leaf pair and ‘stop and go’ phenotype of rg1-1/rg1-1;rg2-2/rg2-3 enabled non- destructive identification of rg1-1/rg1-1;rg2-2/rg2-3 seedlings from the co-segregating wild-type-like rg1-1/rg1-1;RG2/rg2-2 from ~5 dpg onwards which eased comparative monitoring of their development. The growth differences of rg1-1/rg1-1;RG2/rg2-2 between 0% and 3% sucrose media were marginal during the period of ~14 dpg that the seedlings were monitored in the media plates, but were slightly improved on 3% sucrose medium (Figure 2.17A and B). This contrasts with the large differences observed between rg1-1/rg1-1;rg2-2/rg2-3 seedlings grown on 0% versus 3% sucrose media.

2.4.9 Sucrose supplementation restores adult plant height and stature of rg1- 1/rg1-1;rg2-2/rg2-3

To analyze the adult plant phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 seedlings rescued by sucrose supplementation, rg1-1/rg1-1;rg2-2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 seedlings grown on solid 3% sucrose (w/v) MS medium for ~2 weeks were transferred to soil or liquid 3% sucrose (w/v) MS medium. The rg1-1/rg1-1;rg2-2/rg2-3 and rg1-1/rg1- 1;RG2/rg2-2 genotypes were distinguished by the stop-and-go phenotype of rg1-1/rg1- 1;rg2-2/rg2-3 seedlings (described in 2.4.8). For comparison, rg1-1/rg1-1;rg2-2/rg2-3 seedlings grown on 1.5% sucrose (w/v) solid MS medium were transferred to soil at ~2 weeks. The rescued primary meristem of rg1-1/rg1-1;rg2-2/rg2-3 seedlings grown on 3%

73 sucrose medium produced a normal wild-type-like rosette devoid of adventitious shoots (Figure 2.18A). The rg1-1/rg1-1;rg2-2/rg2-3 plants transferred to soil or liquid 3% sucrose medium from solid 3% sucrose medium reached the height of rg1-1/rg1- 1;RG2/rg2-2 (2.18B) and their rosette reached a similar size to that of rg1-1/rg1- 1;RG2/rg2-2 adult plants (2.18C). In comparison to the wild-type-like structure achieved by these rg1-1/rg1-1;rg2-2/rg2-3 plants, the rg1-1/rg1-1;rg2-2/rg2-3 plants grown on 1.5% sucrose medium and transferred to soil showed only partial rescue phenotypes (i.e. retarded seedling growth, failure of primary shoot development followed by development of adventitious shoots resulting in a short adult plant with a compact rosette) (Figures 2.18A-C). However, the silique phenotype of rg1-1/rg1-1;rg2-2/rg2-3 was not rescued by 3% sucrose supplementation. These siliques were small and rarely contained developed seeds, similar to siliques of rg1-1/rg1-1;rg2-2/rg2-3 transferred to soil from 1.5% sucrose medium (2.18B and D).

It is noteworthy that the degree of phenotypic rescue of rg1-1/rg1-1;rg2-2/rg2-3 plants by sucrose supplementation was dependent upon environmental conditions. As described above, when the experiments were conducted under regular growth conditions (i.e. continuous light ~90-120 µmolm-2s-1, relative humidity ~50% and temperature ~22ºC) 3% sucrose supplementation yielded rg1-1/rg1-1;rg2-2/rg2-3 plants with wild-type-like stature. However, when comparative rescue experiments were conducted in sub-optimal growth conditions (i.e. continuous light ~40-50µmolm-2s-1, relative humidity 65%-70% and temperature 23ºC-24ºC - a combination of factors of lower light, higher humidity and higher temperature compared to regular growth conditions), 3% sucrose supplementation could only partially rescue rg1-1/rg1-1;rg2-2/rg2-3 seedlings. These partial rescue phenotypes were similar to rg1-1/rg1-1;rg2-2/rg2-3 grown on 1.5% and 2% sucrose MS media and in soil as described previously where seedling size was small, the primary shoot meristem of rg1-1/rg1-1;rg2-2/rg2-3 ceased to develop and the adult plant resulted from adventitious shoots (examples of partial rescue phenotypes of rg1-1/rg1-1;rg2- 2/rg2-3 are as shown in Figures 2.5-2.11). Under these sub-optimal growth conditions,

74 rg1-1/rg1-1;RG2/rg2-2 seedlings grown on 3% sucrose medium were only marginally retarded compared to regular growth conditions.

2.4.10 Sucrose supplementation restores cell numbers and normal root morphology of rg1-1/rg1-1;rg2-2/rg2-3

To analyze the cellular basis of sucrose rescue in rg1-1/rg1-1;rg2-2/rg2-3 roots, confocal microscopy of propidium iodide or FM4-64 stained roots was performed on rg1-1/rg1- 1;rg2-2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 seedlings grown on 0% and 3% sucrose (w/v) MS media. The ages of the seedlings used for the analyses were 6 dpg for propidium iodide staining and 8 dpg for FM4-64 staining. The rg1-1/rg1-1;rg2-2/rg2-3 seedlings on 0% sucrose MS medium were distinguished from rg1-1/rg1-1;RG2/rg2-2 by the size and morphology differences. The rg1-1/rg1-1;rg2-2/rg2-3 seedlings on 3% sucrose MS medium were distinguished from rg1-1/rg1-1;RG2/rg2-2 by the ‘stop-and-go’ phenotype of rg1-1/rg1-1;rg2-2/rg2-3 seedlings as described in 2.4.8. The compact meristematic zone consisting of fewer cell numbers in rg1-1/rg1-1;rg2-2/rg2-3 on 0% sucrose medium (Figure 2.19A and E) was rescued in rg1-1/rg1-1;rg2-2/rg2-3 on 3% sucrose medium (Figure 2.19B and F) to resemble rg1-1/rg1-1;RG2/rg2-2 (Figure 2.19C, D, G and H). Similarly the overall root tip morphology (i.e. size, shape, cellular arrangement) of rg1- 1/rg1-1;rg2-2/rg2-3 roots was rescued by 3% sucrose supplementation (Figure 2.19A, B, E and F). The cell numbers and root tip morphology were similar between rg1-1/rg1- 1;RG2/rg2-2 grown on 0% and 3% sucrose media (Figure 2.19C, D, G, H).

2.4.11 Sucrose supplementation rescues cell division in the shoot and root apices of rg1-1/rg1-1;rg2-2/rg2-3 seedlings

To test cell division as a basis for phenotypic rescue in the shoot and root apices of rg1- 1/rg1-1;rg2-2/rg2-3 seedlings, transgenic rg1-1/rg1-1;RG2/rg2-2 and rg1-1/rg1-1;rg2- 2/rg2-3 plants containing the mitotic marker construct CycB1;1:GUS (Colon-Carmona et

75 al., 1999) were grown on 0% and 3% sucrose (w/v) MS medium and assayed for β- Glucuronidase activity. CycB1;1:GUS expression evaluated by β-Glucuronidase (GUS) indicates regions with cell division activity.

The rg1-1/rg1-1;rg2-2/rg2-3 seedlings on 0% sucrose MS medium were distinguished from rg1-1/rg1-1;RG2/rg2-2 by size and morphology differences. The 6 dpg rg1-1/rg1- 1;rg2-2/rg2-3 seedlings on 3% sucrose MS medium were distinguished from rg1-1/rg1- 1;RG2/rg2-2 by the ‘stop-and-go’ phenotype of rg1-1/rg1-1;rg2-2/rg2-3 seedlings as described in 2.4.8. The 3 dpg rg1-1/rg1-1;rg2-2/rg2-3 seedlings on 3% sucrose could not be distinguished from the co-segregating rg1-1/rg1-1;RG2/rg2-2 seedlings because the ‘stop-and-go’ phenotype described in 2.4.8 was distinctly observable in rg1-1/rg1-1;rg2- 2/rg2-3 seedlings on 3% sucrose MS medium only after ~5 dpg. Therefore these 3 dpg seedlings were observed as a collection containing both rg1-1/rg1-1;rg2-2/rg2-3 and rg1- 1/rg1-1;RG2/rg2-2 genotypes.

The rg1-1/rg1-1;rg2-2/rg2-3 + CycB1;1:GUS grown on 0% sucrose medium showed CycB1;1:GUS expression (i.e. cell division activity) in a compact region of the shoot apex consistent with its retarded shoot apex development (Figure 2.20A). In comparison, the rg1-1/rg1-1;rg2-2/rg2-3 + CycB1;1:GUS seedlings grown on 3% sucrose medium showed CycB1;1:GUS expression in a larger SAM and young true leaves (Figure 2.20B). Among the seedlings observed on 3% sucrose medium, none were observed to resemble the weak GUS staining and shoot apex morphology of the developmentally retarded rg1- 1/rg1-1;rg2-2/rg2-3 + CycB1;1:GUS seedling shown in Figure 2.20A. Instead, all the seedlings observed on 3% sucrose medium (representative seedlings are shown in Figure 2.20C) showed resemblance in the GUS staining pattern and shoot apex morphology to the rg1-1/rg1-1;RG2/rg2-2 + CycB1;1:GUS shown in Figure 2.20B. In the roots, consistent with the rescued primary root size and morphology of rg1-1/rg1-1;rg2-2/rg2-3 + CycB1;1:GUS by 3% sucrose supplementation, these roots showed a similar CycB1;1:GUS expression pattern at the root tip to rg1-1/rg1-1;RG2/rg2-2 +

76 CycB1;1:GUS grown on 0% and 3% sucrose media (Figures 2.21B-D). In contrast, GUS staining was barely detectable in the rg1-1/rg1-1;rg2-2/rg2-3 grown on 0% sucrose medium that failed to undergo seedling establishment (Figure 2.21A).

2.5 Discussion

Phenotypic characterization of the RanGAP knockdown mutants in this study demonstrates that RanGAPs are required for shoot and root apical integrity, and vegetative plant development in Arabidopsis thaliana. Similar to the yeast and animal systems, requirement of RanGAP for mitotic cell division was shown for plants as well. By complementation assays, redundancy of AtRanGAP1 and AtRanGAP2 was shown for the underlying function. Sucrose rescue experiments suggest the possibility that Arabidopsis RanGAPs may be involved in G1/S or G2/M transitions. However, in yeast and animals RanGAP deficiency has been reported to associate with M-phase defects such as aberrant spindle assembly, chromosome alignment and segregation (described in 2.2.1).

2.5.1 RanGAP, the cell cycle and vegetative plant development

When RanGAP deficient rg1-1/rg1-1;rg2-2/rg2-3 were grown on 1.5% or 2% sucrose media (similar phenotypes were obtained on 1.5% or 2% sucrose media previously referred to as ‘partial rescue phenotypes’), they showed phenotypes that are common with other reported cell-cycle mutants (described in 2.2.5). These phenotypes in rg1- 1/rg1-1;rg2-2/rg2-3 seedlings include smaller seedlings with a compact shoot and short root, retarded growth, variability in seedling phenotypes and lethality of the most severely affected seedlings. In the shoot, delayed primary shoot development with defective true leaf initiation was followed by termination of primary shoot growth and multiple shoot formation from adventitious shoot primordia. In the roots, short meristematic and elongation zones with a drastic reduction in the elongation zone were

77 observed. Root hair development was prominent and occurred closer to the root tip. Disorganized cell files were observed in the root tip. In the adult plants, stunted and bushy plant habit constituted a phenotype common with several reported cell-cycle mutants (see section 2.2.5).

In line with the compact size of rg1-1/rg1-1;rg2-2/rg2-3 vegetative plants obtained on 0%, 1.5% and 2% sucrose media, shoot and root apices were reduced in size compared to rg1-1/rg1-1;RG2/rg2-2. Analysis of the primary root apical meristem (RAM) showed compact size and fewer cells. Concomittently, expression of the mitotic marker CycB1;1::GUS was restricted to compact areas of the shoot and root. Thus, the smaller size of the vegetative plant correlated with reduced size, cell numbers and mitotic activity in the primary meristems indicating defective cell division in the primary meristems as a basis for retarded shoot and root growth leading to compact plants. Possible cellular and molecular basis for the developmental phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 are discussed in sections 2.5.2 and 2.5.3 below.

The phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 that resemble other cell-cycle mutants together with the preliminary cellular and molecular data presented here are consistent with RanGAP functioning as a positive regulator for cell division in the apical vegetative meristems and thereby in undifferentiated vegetative growth. The absence of these cell cycle-related phenotypes in the RanGAP single mutants and the finding that either RanGAP1 or RanGAP2 could rescue these phenotypes in rg1-1/rg1-1;rg2-2/rg2-3 show that the two RanGAPs are redundant for the underlying molecular function(s). The finding that localization of the GFP-N7 nuclear import marker remained unaffected in the root-cap and non-root cap regions in rg1-1/rg1-1;rg2-2/rg2-3 together with similar results in the inducible RanGAP RNAi lines provide a preliminary indication that the root phenotypes described may be caused by nuclear transport-independent functions of RanGAP.

78 2.5.2 RanGAP in shoot growth and development

Pertubation of shoot apical growth in rg1-1/rg1-1;rg2-2/rg2-3 is indicated by the small shoot apex size and retarded growth with limited initiation of true leaves and their slow development. Pertubation of integrity of the shoot apex development is indicated by disrupted true leaf initiation and random development of adventitious shoots in all regions of the meristem including the central zone. Thus, the two functions attributed to the central zone of the primary shoot apical meristem (SAM); self-renewal and regulation of meristem integrity (Steeves and Sussex, 1989; Laux et al., 1996) seem to be affected in the rg1-1/rg1-1;rg2-2/rg2-3 mutant suggesting that RanGAP plays a role in these functions. The primary shoot that is initiated by the activity of SAM halts development in rg1-1/rg1-1;rg2-2/rg2-3 and the adult plant develops from the adventitious shoots that result from the activity of the adventitious meristems initiated at the shoot apex.

The SAM plays a critical role in the initiation, growth and development of leaf primordia including establishment of dorsiventrality (Van Lijsebettens and Clarke, 1998). Therefore a defective SAM function could explain the defects in true leaf initiation and their development observed in rg1-1/rg1-1;rg2-2/rg2-3. The shoot apex development in rg1- 1/rg1-1;rg2-2/rg2-3 also suggests a relationship between SAM function and adventitious shoot primordia development. When the primary SAM function was defective in plants grown on 1.5% and 2% sucrose media and in soil, adventitious shoot primordia were initiated randomly at the shoot apex. However, when cell division and the primary shoot growth were restored by 3% sucrose supplementation, adventitious primordia initiation was suppressed and the adult plant developed from the activity of the primary SAM, as in rg1-1/rg1-1;RG2/rg2-2. Supporting a hypothesis that initiation of adventitious shoots results from improper functioning of the primary SAM, it has been previously shown that mechanical destruction of the central zone leads to adventitious shoot meristems in the periphery of the shoot apex (Loiseau, 1959).

79 Several adventitious shoots that develop into small rosettes and short inflorescences in rg1-1/rg1-1;rg2-2/rg2-3, together with the excessive branching of inflorescences account for the short and bushy habit on 1.5% and 2% sucrose media and in soil. Excessive branching is reported to occur as a result of infertility (Robinson-Beers et al., 1992, Hensel et al., 1994). When rg1-1/rg1-1;rg2-2/rg2-3 were grown for up to ~2 weeks on 3% sucrose plates and were then transferred either to soil or liquid 3% sucrose medium, the primary SAM produced a vegetative rosette similar in size to rg1-1/rg1-1;RG2/rg2-2 plants. The inflorescences were restored to resemble the height and stature of rg1-1/rg1- 1;RG2/rg2-2 when observed at ~55 dpg. In contrast, when rg1-1/rg1-1;rg2-2/rg2-3 plants were transferred to soil from 1.5% or 2% sucrose media, or when they were grown in soil from seed stage they showed a dwarf habit. This could be due to the inherent differences in the development of adventitious shoots versus the primary shoot, respectively. Supporting this, a dwarf plant habit has been associated with mutants that form the adult plant from the development of adventitious shoots (Bundock and Hooykaas, 2002; Binarova et al., 2006; Wang and Liu, 2006; Xia et al., 2006; ).

Given the central function of WUS in primary SAM development, it appears worthwhile to consider the multiple shoot formation commonly seen in cell cycle mutants with respect to WUS expression. WUS expression has been analyzed in some of the cell cycle mutants that are reported to give rise to multiple shoots. Among them, examples for both 1) WUS underexpression (e.g. AtCDC5 mutant) (Lin et al., 2007) and 2) WUS overexpression (e.g. INCURVATA 2 mutant) (Barrero et al., 2007) or an enlarged WUS expression domain (e.g. BRU 1 and FASCIATA mutants) (Mayer et al., 1998; Kaya et al., 2001; Takeda et al., 2004) have been reported. Both scenarios could potentially disrupt the proper shoot meristem function; 1) WUS underexpression by reduced cell division and, 2) WUS overexpression or an enlarged WUS expression domain by over- proliferation of stem cells. Further, both these scenarios could trigger the development of adventitious and/or lateral shoot primordia as a result of the loss of proper function of the primary SAM. However, phenotypes specific to WUS underexpression or WUS

80 overexpression/misexpression seem to manifest in the mutants belonging to the respective category exclusively. For example, fasciation of stems and organs were observed in the mutants with dispersed expression of WUS mRNA such as bru1, fas1 and fas2 (Mayer et al., 1998; Kaya et al., 2001; Takeda et al., 2004) but these phenotypes were not reported in the WUS underexpressing mutants (Laux et al., 1996; Lin et al., 2007).

The shoot phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 resemble wus-1 phenotypes including reduced cell division, non-fasciation of stems, floral organs and siliques, adventitious shoot primordia initiation in the shoot apex including in the central zone (Laux et al., 1996). Further, the adventitious rosettes and inflorescences were able to develop with normal phyllotaxy in both rg1-1/rg1-1;rg2-2/rg2-3 and wus-1 mutant. Given the observation that RanGAP is required for primary shoot development (which is dependant on the activity of the primary SAM), it is intriguing how the adventitious rosettes and inflorescences (dependent on the activity of adventitious meristems) were able to develop with normal phyllotaxy. It was proposed in the case of wus-1 mutant that the diminishing importance of WUS as a positive regulator of cell division in the shoot as plant development progresses may allow development of adventitious shoots with normal phyllotaxy later on (Laux et al., 1996; Gallois et al., 2002). A similar scenario could be true for rg1-1/rg1-1;rg2-2/rg2-3 where the importance of RanGAP for cell division in adventitious shoot meristem is diminished, either through the WUS pathway or an independent pathway. It is noteworthy that the defects in floral morphology in the wus-1 mutant that are possibly specific to WUS depletion in the floral apex were not observed under RanGAP deficiency.

2.5.3 RanGAP in root growth and development

At the level of depletion of RanGAPs in rg1-1/rg1-1;rg2-2/rg2-3, RanGAPs seem dispensable for the establishment of the root apical meristem (RAM) but essential for its

81 maintenance. This is similar to the scenario observed with the primary SAM where the developmental defects were observed during post-embryonic development. Root growth phenotypes (i.e. short roots, retarded growth and termination of root growth in severe manifestations) are likely a direct result of the limited/halted supply of cell numbers resulting from cell division defects in the root apex that limited the spans of meristematic, elongation and differentiation zones.

Root development phenotypes observed in rg1-1/rg1-1;rg2-2/rg2-3 include placement of the root hair-zone closer to the root apex, more prominent root hair development and disorganization cells files. These phenotypes were commonly reported in other cell cycle mutants as well (described in 2.2.5). Shortened meristem and elongation zones could underlie the placement of root hair-containing differentiated cells closer to the root apex. Alternatively, an altered signaling mechanism could contribute to the development of root hairs closer to the root apex. Regulation of exit from cell cycle to enter endoreduplication or other mechanisms that underlie enlargement/elongation of a cell is fine-tuned (Cebolla et al., 1999; De Veylder et al., 2001; Inze, 2005; Menges et al., 2006) which may be true for differentiation as well. These processes could be expedited in the cell division mutants including rg1-1/rg1-1;rg2-2/rg2-3. GLABRA2 (GL2), encoding a homeodomain transcription factor required for repression of root hair formation (Ohashi et al., 2003) has shown to be down-regulated in the mutant of NUCLEOSOME ASSEMBLY PROTEIN1 (NAP1)-RELATED PROTEIN (NRP) which is defective in cell cycle progression. NRP encodes a histone chaperone required during DNA replication (Zhu et al., 2006).

Misexpression of the SCARECROW transcription factor in the root has been shown in two of the cell cycle mutants; fas and teb that show disorganized cell files (Kaya et al., 2001; Inagaki et al., 2006). Periclinal division of cortex/endodermal initial cells, a function known for the SCARECROW (SCR) gene, is postulated to underlie establishment of disorganized cell files in these mutants (Kaya et al., 2001; Inagaki et al.,

82 2006). A similar phenomenon could hold true for rg1-1/rg1-1;rg2-2/rg2-3 as well, leading to disorganized cell files at the root tip. Also, the requirement of SCR expression for maintenance of quiescent center and stem cell niche (Sabatini et al., 2003) could be causal for short roots in the cell cycle mutants including rg1-1/rg1-1;rg2-2/rg2-3. The abundant obliquely placed cell walls, misshaped cells and cell wall stubs reported in the ground tissue of roots of inducible RanGAP RNAi lines (Xu et al., 2008) are likely a more profound manifestation of the cellular defects observed in the root cap region of rg1-1/rg1-1;rg2-2/rg2-3.

2.5.4 Sucrose rescue of RanGAP mutant phenotypes

Sucrose supplementation (i.e. 1.5%, 2% or 3% sucrose) or withdrawal (i.e. 0% sucrose) caused major changes in rg1-1/rg1-1;rg2-2/rg2-3 vegetative growth and development compared to the marginal changes observed in rg1-1/rg1-1;RG2/rg2-2 plants. Thus at the whole plant level, seedling size and morphology as well as adult plant height and stature were rescued with increased sucrose supplementation. In the vegetative apices, shoot and root apex sizes and morphology, size and cell numbers in the RAM, and expression of the mitotic marker CycB1;1::GUS were rescued by sucrose. These results together with the knowledge on involvement of sucrose in cell cycle progression (described in 2.2.4) and the resemblance of rg1-1/rg1-1;rg2-2/rg2-3 to cell cycle mutants when grown on 1.5% or 2% sucrose media suggest that supplementation of sucrose causes phenotypic rescue likely via rescue of cell division in the apical meristems.

Sucrose has shown to be a positive regulator of cell division by causing Cyclin D induction in G1/S transition (see section 2.2.4). One possibility for the rg1-1/rg1-1;rg2- 2/rg2-3 phenotype is thus that the mutants are deficient in G1/S cell cycle commitment upstream of cyclin D induction causing cell cycle perturbation-like phenotypes, which are rescued by sucrose supplementation that induce cyclin D. Assays of cyclin D levels and stage of cell cycle arrest in the vegetative meristems would be necessary to test this

83 hypothesis. If shown, RanGAP could be involved in the G1/S molecular mechanism directly or via an indirect mechanism. One possibility of direct involvement in G1/S transition may be via involment of RanGAP in the cyclin D induction pathways downstream of hormonal signaling such as cytokinin, auxin and brassinosteroids (described in 2.2.4). Alternatively, a mechanism in a different stage of the cell cycle may be possible for this effect, such as in the case of the stip mutant (described in 2.2.4).

The silique phenotype of rg1-1/rg1-1;rg2-2/rg2-3 was not rescued by 3% sucrose supplementation, under conditions that rescued the vegetative phenotypes. This could mean that the underlying molecular mechanism of the cell division defect in vegetative development may not be the cause for female gametophyte arrest during mitotic phases in the RanGAP deficient mutants as described in Chapter 1 and Rodrigo-Peiris et al., 2011. It may also be possible that the supplement of sucrose provided through the growth medium is not sufficiently transported to the developing gametophyte to cause the rescue.

While near complete rescue of rg1-1/rg1-1;rg2-2/rg2-3 phenotypes were achieved by 3% sucrose supplementation under regular growth conditions, profound defects in development amounting to only partial rescue of vegetative phenotypes were achieved under sub-optimal growth conditions despite supply of 3% sucrose. Thus, growth and developmental phenotypes likely pertaining to defective cell division in the apical meristems manifested in rg1-1/rg1-1;rg2-2/rg2-3 under these sub-optimal conditions on 3% sucrose supplementation. In comparison, only mild growth defects were caused in the rg1-1/rg1-1;RG2/rg2-2 plants in these sub-optimal growth conditions compared to the regular growth conditions. Thus the presence of RanGAP in rg1-1/rg1-1;RG2/rg2-2 appears to provide a buffering effect to adverse environmental effects in terms of cell cycle progression. In plants, it is known that cell cycle regulation responds to stimuli from both the internal and external environment (Richard et al., 2002). The sensitivity of rg1-1/rg1-1;rg2-2/rg2-3 to the stipulated sub-optimal conditions compared to rg1-1/rg1- 1;RG2/rg2-2 leading to more profound cell division-related defects may mean that the rg

84 mutation is sensitive to the environmental factor(s). If so, this could place RanGAP in the signaling cascade downstream of these environmental factor(s) leading to cell cycle activation (Medford et al., 1992). Another possibility is that a compensatory cell division mechanism involving RanGAP in the rg1-1/rg1-1;RG2/rg2-2 may account for the buffering effect in rg1-1/rg1-1;RG2/rg2-2, which is deficient in rg1-1/rg1-1;rg2-2/rg2-3.

Despite the proposed importance of sugars as a carbon source and signaling molecules for cell division and development in the meristems, only a few mutants have been identified to link sugar to meristem function. The stip mutation leading to cell division defects and meristem function via derepression of the cell cycle negative regulator TSS, and the rescue of these phenotypes by sucrose supplementation that leads to transcriptional repression of TSS is the most studied example (Wu et al., 2005; Wu et al., 2007; Skylar et al., 2010a; Skylar et al., 2010b; Skylar et al., 2011). Recently, the characterization of two FANTASTIC FOUR genes FAF2 and FAF4 have shown to regulate shoot meristem size and function by acting via WUS. Their overexpression has shown to arrest shoot and root growth (Wahl et al., 2010). Addition of exogenous sucrose has shown to rescue root growth arrest, however the underlying molecular mechanism remains to be studied (Wahl et al., 2010). ramosa 3, a mutant in trehalose synthesis (trehalose is a non-reducing disachcharide proposed to be involved in a regulatory role in sugar signaling rather than a metabolic role) has shown defects in axillary meristem development in maize inflorescence (reviewed in Eveland and Jackson, 2011). Together with its restricted localization to the base of these meristems, RAMOSA is proposed to link sugar signaling to inflorescence meristem development. Based on the shoot apical growth and development phenotypes observed in rg1-1/rg1-1;rg2-2/rg2-3 in this study and their rescue by sucrose supplementation, it may be possible that RanGAP links sugar signaling to vegetative meristem development via the cell cycle.

85 2.5.5 Future directions

A preliminary assay using the GFP-N7 nuclear import protein indicated that the phenotypic defects in the RanGAP-deficient Arabidopsis mutants studied here are possibly independent of the involvement of RanGAP in nuclear transport. Further and alternative nucleocytoplasmic transport assays are necessary to confirm this finding. Nucleocytoplamic transport assays using GFP-fused shuttling markers that evaluate nuclear import and export concomitantly (similar to the construct used by Kusano et al., 2004) may provide valuable insight.

With the knowledge that sucrose rescues cell cycle arrest at G1/S boundary by cyclin D induction (den Boer and Murray, 2000; Gaudin et al., 2000; Riou-Khamlichi et al., 2000; Wu et al., 2005), one hypothesis remains that rg1-1/rg1-1;rg2-2/rg2-3 is deficient in G1/S commitment. To test this, cyclin D expression in rg1-1/rg1-1;rg2-2/rg2-3 needs to be analyzed, for which in situ hybridization appears a suitable approach (Dewitte et al., 2003). However, particularly due to the recent unexpected finding that the stip mutant showed cell cycle arrests at G2 phase but was rescued by sucrose supplementation (Wu et al., 2005; Wu et al., 2007; Skylar et al., 2011), experiments to evaluate the stage of cell cycle arrest in rg1-1/rg1-1;rg2-2/rg2-3 would be informative. Ploidy-level analysis experiments as performed by Wu et al., 2007 would likely be a feasible approach to determine arrest at pre-DNA replication (i.e. 2C ploidy level) to support a G1 phase arrest.

To understand the molecular basis underlying the RanGAP-knockdown vegetative phenotypes, an enhancer/suppressor screen using the semi-fertile rg1-1/rg1-1;rg2-2/rg2-2 mutant holds potential. Further, a comparative microarray analysis (Zhu et al., 2006) between rg1-1/rg1-1;rg2-2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 grown on 0%, 1.5-2% and 3% sucrose could provide valuable insight in this regard.

86 Due to the similarity of the rg1-1/rg1-1;rg2-2/rg2-3 and the wus-1 mutant in shoot development, it appears worthwhile to test for a functional relationship between RanGAP and WUS. Introgression of wus-1 into rg1-1/rg1-1;rg2-2/rg2-3 mutant background and comparative phenotypic analysis with wus-1 and rg1-1/rg1-1;rg2-2/rg2-3 mutants may shed light whether a functional relationship between WUS and RanGAP exists in SAM development. Given the fact that WUS is a central regulator of SAM, it may be possible that RanGAP could in fact regulate cell division via WUS itself. Analysis of WUS expression and subcellular localization in the shoot apex of rg1-1/rg1-1;rg2-2/rg2-3, and rescue experiments via WUS expression in rg1-1/rg1-1;rg2-2/rg2-3 could provide useful information in this regard.

Based on the phenotypic similarity between rg1-1/rg1-1;rg2-2/rg2-3, other cell cycle mutants that are defective in different stages of cell division and wild-type plants treated chemically or by radiation to perturb the cell cycle, I have proposed manifestation of overlapping phenotypic defects in plant vegetative development which could be general downstream effects of an aberrant cell cycle. The understanding of these common phenotypes could facilitate rapid identification of more mutants defective in various stages of the cell cycle such as by large-scale mutant screens, which could enhance our understanding of the genetic and molecular basis of the cell cycle. With such extension of cell cycle mutants from the sparse collection existing currently and with more focused and detailed characterization of each of these mutants, it may be possible not only to confirm this hypothesis (i.e. that common downstream effects and phenotypes exist resulting from an aberrant cell cycle) but also to identify phenotypes and thereby molecular players that are restricted to certain phases of the cell cycle. Particularly, sucrose rescue assay and further experimentation on these cell cycle mutants could lead to novel findings that connect sugar signaling and the cell cycle. By evaluating the expression of the known genes in root and shoot pattern formation (e,g, SCR, WUS), genes involved in cell differentiation, plant growth and development (e.g. PLT2, GL2) and stress response genes [e.g. PARPs (Vanderauwera et al., 2007), Peroxidases (Zhu et

87 al., 2006), Cytochrome P450 family proteins (Zhu et al., 2006), Universal stress protein (USP) family proteins (Zhu et al., 2006)] in these mutants or by general microarray analyses (Zhu et al., 2006), it may be possible to elucidate molecular pathways relating cell division cycle to cell differentiation, pattern formation, plant growth, development and stress response mechanisms.

88 A rg2-2 200bp

ATG

Promoter Ex In Ex

5’ UTR ORF 3’ UTR

WT (Col) rg2-2 70 kD α -RanGAP2

α -MFP1

Figure 2.1: T-DNA insert-position and protein expression of rg2-2 allele. (A) Schematic representation of RanGAP2 gene structure and the position of T-DNA insert in the rg2-2 allele. ORF, open reading frame; Ex, Exon; In, Intron; ATG, open reading frame start position; blocked arrow, open reading frame end position. Triangle depicts the position of T-DNA insertion. Scale bar: 200 bp (B) Immunoblot analysis shows that rg2- 2 is a knockdown allele. Total protein extracts from 8 day-old Arabidopsis seedlings of Columbia wild-type [WT (Col)] and rg1-1/rg1-1;rg2-2/rg2-2 mutant (Col) were incubated with anti-RanGAP2 antibody. As the loading control, the blot was stripped and re-incubated with anti-MFP1 antibody.

Figure 2.2: Seedling and adult plant phenotypes of the RanGAP knockdown rg1- 1/rg1-1;rg2-2/rg2-2 mutant. (A) 5 dpg seedlings show mild growth and development phenotypes in the rg1-1/rg1-1;rg2-2/rg2-2 mutant compared to wild-type. Smaller size of the mutant is seen. First true leaves are visible in the wild-type but not visible in the mutant. (B) Comparative adult plants at ~6 weeks post-germination. Both plants show similar vegetative morphology and adult height. The mutant shows a short silique phenotype. Seedlings in (A) were grown on 2% sucrose (w/v) MS medium. Plants in (B) were grown in soil. Details of growth conditions are specified in Materials and methods.

90 Figure 2.2

A rg1-1/rg1-1; Wild-type (Col) rg2-2/rg2-2

B rg1-1/rg1-1; Wild-type (Col) rg2-2/rg2-2

Figure 2.3: Reproductive phenotypes of the RanGAP knockdown rg1-1/rg1-1;rg2- 2/rg2-2 mutant and phenotypic complementation. (A) A short silique of rg1-1/rg1- 1;rg2-2/rg2-2 is shown compared to wild-type (Col). Less seeds are developed in the rg1-1/rg1-1;rg2-2/rg2-2 silique, and many developmentally arrested ovules are seen as white and shrunken miniscule structures (example shown by arrow). (B)-(G) Ovules of rg1-1/rg1-1;rg2-2/rg2-2 pistils from floral stage 12-late (floral stages as defined in Table 1.2) showing variable developmental phenotypes in the female gametophytes and integuments. Ovules were cleared with chloral hydrate solution and observed with DIC microscopy as described in 1.3.5. Ovules are arrested in a range of gametophyte development stages from pre-meiosis to post-mitotic stages; (B) megaspore mother cell (C) tetrad (D) 1-nuclear embryo sac (E) 2-nuclear embryo sac (F) 4-nuclear embryo sac and (G) 8-nuclear embryo sac (only 3 nuclei are visible in the presented plane). Nuclei are encircled. Integument development defects (i.e. shorter integuments) are seen in (F) and (G). Comparative wild-type (Col) ovules at the respective developmental stages are presented in Figure 1.5. Scale bars: 50 µm. (H) Silique lengths of rg1-1/rg1-1;rg2-2/rg2- 2 plants carrying RanGAP1 promoter-driven RanGAP1 construct [rg1-1/rg1-1;rg2-2/rg2- 2 + RG1] (Extreme right) are rescued to resemble silique lengths of wild-type (Col) (Extreme left). Short siliques of rg1-1/rg1-1;rg2-2/rg2-2 plants are shown in the middle. (I) Comparative immunoblot analysis of siliques from the genotypes shown in (H). Total proteins were extracted from a pool of whole siliques and detected using anti-RanGAP1 antibody. Siliques with rescued size from rg1-1/rg1-1;rg2-2/rg2-2 + RG1 plants (lines 1, 2 and 3) and siliques of wild-type (Col) show RanGAP1 protein expression. rg1-1/rg1- 1;rg2-2/rg2-2 (negative control) lacks RanGAP1 protein expression. The rg1-1/rg1- 1;rg2-2/rg2-2 + RG1 siliques used for the analysis are fruits of T1 plants upon transformation of T0 rg1-1/rg1-1;rg2-2/rg2-2 with the RanGAP1 promoter-driven RanGAP1 construct. Coomassie brilliant blue staining of a replica gel is shown at the bottom as the loading control.

92 Figure 2.3

rg1-1/rg1-1;rg2-2/rg2-2

WT (Col)

B C D

E F G

H rg1-1/rg1-1; rg1-1/rg1-1; WT (Col) rg2-2/rg2-2 rg2-2/rg2-2 + RG1

Figure 2.4: Embryo observations and molecular characterization of rg1-1/rg1-1;rg2- 2/rg2-3 mutant. (A) A representative mature embryo resulting from the cross of ♀ rg1- 1/rg1-1;rg2-2/rg2-2 X ♂ rg1-1/rg1-1;RG2/rg2-3 observed with DIC light microscopy. rg1-1/rg1-1;RG2/rg2-2 and rg1-1/rg1-1;rg2-2/rg2-3 embryos were not distinguishable due to the absence of discernable phenotypes under the resolution obtained. Scale bars: 50 µm. (B) A representative genotyping result that distinguish between rg1-1/rg1- 1;RG2/rg2-2 and rg1-1/rg1-1;rg2-2/rg2-3 seedlings. Arrows indicate the expected position for the PCR products of RG2 and rg2-3 alleles. In rg1-1/rg1-1;RG2/rg2-2, presence of the PCR product for RG2 and absence of the PCR product for rg2-3 is seen. In rg1-1/rg1-1; rg2-2/rg2-3, presence of the PCR product for rg2-3 and absence of the PCR product for RG2 is seen. Genotyping methodology is as described in 2.3.2. (C) Immunoblot analysis shows that rg1-1/rg1-1;rg2-2/rg2-3 mutant contains a further knockdown level of RanGAP2 compared to rg1-1/rg1-1;rg2-2/rg2-2 mutant. Total protein extracts from several 12 day-old Arabidopsis seedlings of the two mutants in a similar Columbia/Wassilkja hybrid background were incubated with anti-RanGAP2 antibody. Coomassie brilliant blue staining of a replica gel is shown at the bottom as the loading control.

94 Figure 2.4

Shoot Root apex apex

rg1-1/rg1-1;RG2/rg2-2 or rg1-1/rg1-1;rg2-2/rg2-3 embryo

rg1-1/rg1-1;RG2/rg2-2 rg1-1/rg1-1;rg2-2/rg2-3

RG2 rg2-3 RG2 rg2-3 RG2 rg2-3 RG2 rg2-3 RG2 rg2-3 RG2 rg2-3 RG2 rg2-3 RG2 rg2-3 RG2 RG2 rg2-3 rg2-3

Plant 1 Plant 2 Plant 3 Plant 4 Plant 1 Plant 3 Plant 4 Plant 2

rg1-1/rg1-1; rg1-1/rg1-1; rg2-2/rg2-3 rg2-2/rg2-2

α -RanGAP2

Coomassie

Figure 2.5: Seedling phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 mutant. (A) rg1-1/rg1- 1;rg2-2/rg2-3 show seedling phenotypes compared to rg1-1/rg1-1;RG2/rg2-2. 5 dpg seedlings were photographed. Smaller size of rg1-1/rg1-1;rg2-2/rg2-3 seedlings with less expanded cotyledons and shorter roots are seen. Black horizontal lines indicate the position of the root tips at 4 dpg (B) Scanning electron micrographs of 2 dpg seedlings showing smaller size and delayed development of shoot apex in rg1-1/rg1-1;rg2-2/rg2-3. Cotyledons are less expanded in rg1-1/rg1-1;rg2-2/rg2-3 seedlings. The first true leaves are visible in rg1-1/rg1-1;RG2/rg2-2 but not in rg1-1/rg1-1;rg2-2/rg2-3. Scale bars: 50 µm. Seedlings were grown on 2% sucrose (w/v) MS medium.

96 Figure 2.5

rg1-1/rg1-1; rg1-1/rg1-1; rg1-1/rg1-1; rg1-1/rg1-1; rg2-2/rg2-3 RG2/rg2-2 rg2-2/rg2-3 RG2/rg2-2

rg1-1/rg1-1;RG2/rg2-2 rg1-1/rg1-1;rg2-2/rg2-3

rg1-1/rg1-1;rg2-2/rg2-3 2 dpg rg1-1/rg1-1;RG2/rg2-2 2 dpg A B

C rg1-1/rg1-1;rg2-2/rg2-3 4 dpg D rg1-1/rg1-1;RG2/rg2-2 4 dpg

Figure 2.6: Scanning electron micrographs show delayed shoot apex development in rg1-1/rg1-1;rg2-2/rg2-3 seedlings. Shoot apices of (A, B) 2 dpg and (C, D) 4 dpg seedlings of (A, C) rg1-1/rg1-1;rg2-2/rg2-3 and (B, D) rg1-1/rg1-1;RG2/rg2-2 are shown. (A) True leaves are not initiated in rg1-1/rg1-1;rg2-2/rg2-3 at 2 dpg. (B) First pair of true leaves are initiated in rg1-1/rg1-1;RG2/rg2-2 at 2 dpg. (C) One of the emerging first true leaves of rg1-1/rg1-1;rg2-2/rg2-3 at 4 dpg is seen as a cylindrical protrusion (arrow). The smaller second leaf protrusion is hidden in this view. (D) Shoot apex of rg1-1/rg1-1;RG2/rg2-2 is at the development stage of four true leaves at 4 dpg. Scale bars: 50 µm. Seedlings were grown on 2% sucrose (w/v) MS medium.

98 A rg1-1/rg1-1; rg2-2/rg2-3

B rg1-1/rg1-1; C rg2-2/rg2-3

rg1-1/rg1-1; rg2-2/rg2-3

D rg1-1/rg1-1; E rg1-1/rg1-1; rg2-2/rg2-3 RG2/rg2-2

Figure 2.7: Development of the first pair of true leaves is abnormal in rg1-1/rg1- 1;rg2-2/rg2-3 seedlings and successive initiation of true leaves in a radial pattern is perturbed. (A)-(D) rg1-1/rg1-1;rg2-2/rg2-3 seedlings. (A) The first true leaf pair is terminated at minuscule sizes in a 12 dpg seedling. Arrow in (B) points to a crinkled first true leaf in a 16 dpg seedling. The 12 dpg seedling in (C) contains a bifid leaf (arrow) and a poorly expanded oblong-shaped leaf (arrowhead). Arrow in (D) points to a first leaf in a 9 dpg seedling that is cylindrical and lacks dorso-ventral tissue specialization. The inset shows a cross section of the cylindrical leaf. (E) A rg1-1/rg1-1;RG2/rg2-2 seedling at 12 dpg with the radial organization of true leaves. Scale bars in A-E: 1.25 mm. Scale bar in inset: 30 µm. Seedlings were grown on 2% sucrose (w/v) MS medium.

Figure 2.8: Histological analysis reveals disorganized cell arrangement in the developing true leaves and shoot apex of rg1-1/rg1-1;rg2-2/rg2-3. Longitudinal histological sections of the shoot apex of (A) rg1-1/rg1-1;RG2/rg2-2 and (B) rg1-1/rg1- 1;rg2-3/rg2-3 seedlings at 4 dpg. Vertical sections were obtained through a plane perpendicular to the plane of the cotyledons to reveal the cellular arrangement of the first two true leaves (arrowheads) in the projected view. White arrows in (A) point to stipules. (A) shows uniform cell shapes and layered organization of cells in the first true leaves of rg1-1/rg1-1;RG2/rg2-2. (B) shows irregular cell shapes, sizes and disturbed cell layer organization is the first true leaves of rg1-1/rg1-1;rg2-2/rg2-3. Abundant irregular cell division planes are seen (examples shown by arrows). Shoot apex of rg1-1/rg1-1;rg2- 2/rg2-3 contains a cell mass that resembles an adventitious shoot meristem by gross cellular arrangement (curved line). Note that section (B) represents an anterior tangential plane with respect to the center of the shoot which dissects the adventitious shoot meristem shown. Scale bars: 50 µm. Seedlings were grown on 2% sucrose (w/v) MS medium.

100 Figure 2.8

rg1-1/rg1-1;RG2/rg2-2

rg1-1/rg1-1;rg2-2/rg2-3

101

Figure 2.9: Severely affected rg1-1/rg1-1;rg2-3/rg2-3 seedlings are lethal. Adventitious shoot meristems of surviving rg1-1/rg1-1;rg2-3/rg2-3 seedlings give rise to a compact shoot with multiple rosettes. (A) A lethal rg1-1/rg1-1;rg2-3/rg2-3 seedling at 9 dpg. Arrow points to epidermal gland-like structures. (B) A lethal rg1- 1/rg1-1;rg2-3/rg2-3 seedling at 10 dpg. Arrow points to a callus-like structure. (C) Scanning electron micrograph of the shoot apex of a 12 dpg seedling of rg1-1/rg1-1;rg2- 3/rg2-3. Randomly placed multiple adventitious shoots at the shoot apex are seen. Arrowheads point to the putative first pair of true leaves; Black arrows point to leaves developing from random positions of the shoot apex. Red arrow points to an adventitious shoot primordium. (D) Scanning electron micrograph of the shoot apex of a 12 dpg seedling of rg1-1/rg1-1;RG2/rg2-2. Radial organization of true leaves is seen. Scale bars: 500µm. (E) 29 dpg plants of rg1-1/rg1-1;RG2/rg2-2 (left) and rg1-1/rg1-1;rg2-3/rg2-3 (right). (F) and (G) are enlarged views of rg1-1/rg1-1;rg2-3/rg2-3 plants shown in (E). Red arrows in (F) and (G) point to the positions of the adventitious shoots. Blue arrow in (F) points to a developing inflorescence stem. Seedlings were grown on 2% sucrose (w/v) MS medium. Plants in (E)-(G) were transferred to soil at 13 dpg.

102 Figure 2.9

A B

rg1-1/rg1-1;rg2-2/rg2-3 rg1-1/rg1-1;rg2-2/rg2-3

C D

rg1-1/rg1-1;rg2-2/rg2-3 rg1-1/rg1-1;RG2/rg2-2

E F G

rg1-1/rg1-1; rg1-1/rg1-1; RG2/rg2-2 rg2-2/rg2-3

rg1-1/rg1-1; rg1-1/rg1-1; rg2-2/rg2-3 rg2-2/rg2-3

103

Figure 2.10: Root morphological and anatomical phenotypes of rg1-1/rg1-1;rg2- 2/rg2-3. Primary root of 5 dpg seedlings of (A) rg1-1/rg1-1;rg2-2/rg2-3 and (B) rg1- 1/rg1-1;RG2/rg2-2. Arrowheads demarcate the approximate elongation zone with the upper arrowhead in each root indicating the position of the first observable root hair. Note the prevalent root hairs in (A). Propidium iodide staining of primary roots of 5 dpg (C, E) rg1-1/rg1-1;rg2-2/rg2-3 and (D, F) rg1-1/rg1-1;RG2/rg2-2. Confocal optical sections were obtained through the mid plane in (C) and (D) and in a sub-surface plane in (E) and (F). White arrow in (C) points to the central root cap region containing disorganized cell files. The circle in (C) highlights an area that shows disorganized arrangement of cell files in the ground tissue with short spans of displaced and additional cell files. Black arrow in the circle indicates a short additional cell file. Arrowhead in the circle points to a displaced/misaligned cell file. Arrow in (E) points to the lateral root cap region containing disorganized cell files. Arrowheads in (E) point to oblique cell walls in the lateral root cap region. rg1-1/rg1-1;RG2/rg2-2 shows organized cell files and regular cell shapes in (D) and (E). Magnifications are identical in the comparative pairs (A)/(B), (C)/(D) and (E)/(F). Scale bars: 50 µm. Seedlings were grown on 2% sucrose (w/v) MS medium.

104 Figure 2.10

A B

rg1-1/rg1-1; rg2-2/rg2-3

rg1-1/rg1-1; RG2/rg2-2 C D

rg1-1/rg1-1; rg1-1/rg1-1; rg2-2/rg2-3 RG2/rg2-2 E F

rg1-1/rg1-1; rg1-1/rg1-1; rg2-2/rg2-3 RG2/rg2-2

105 rg1-1/rg1-1; A B C rg2-2/rg2-3

rg1-1/rg1-1; rg2-2/rg2-3

rg1-1/rg1-1; rg1-1/rg1-1; rg1-1/rg1-1; rg2-2/rg2-3 RG2/rg2-2 rg2-2/rg2-3

E rg1-1/rg1-1; rg2-2/rg2-3

rg1-1/rg1-1;RG2/rg2-2 rg1-1/rg1-1; rg1-1/rg1-1; RG2/rg2-2 rg2-2/rg2-3

Figure 2.11: Adult plant phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 mutant. (A) 40 dpg rg1-1/rg1-1;rg2-2/rg2-3 plants show smaller rosette size and shorter height compared to rg1-1/rg1-1;RG2/rg2-2. (B) and (C) are enlarged views of rg1-1/rg1-1;rg2-2/rg2-3 plants shown in (A). (D) 65 dpg rg1-1/rg1-1;rg2-2/rg2-3 shows a short and bushy stature of mature plant compared to rg1-1/rg1-1;RG2/rg2-2. Shorter inflorescence stems of rg1- 1/rg1-1;rg2-2/rg2-3 are branched excessively. rg1-1/rg1-1;RG2/rg2-2 has terminated flowering/fruiting and is senescing while rg1-1/rg1-1;rg2-2/rg2-3 is in the flowering/ fruiting stage. (E) Shorter siliques of rg1-1/rg1-1;rg2-2/rg2-3 compared to rg1-1/rg1- 1;RG2/rg2-2. Seedlings were grown on 2% sucrose (w/v) MS medium and transferred to soil at 13 dpg.

106 A

Juvenile Adult Petiole Stem Shoot Hypocotyl leaf leaf apex RanGAP1

RanGAP2

9.0 10.4 >11.79

Root hair Elongation Meristematic zone zone zone RanGAP1

RanGAP2

9.0 10.54 >12.87

Figure 2.12: Microarray expression data shows enhanced RanGAP expression in the shoot apex and root meristem. Microarray expression data were retrieved from GENEVESTIGATOR (Zimmermann et al., 2004). The intensity of the yellow scale corresponds to the mean log2 normalized expression values of RanGAPs on the microarrays, as shown in the key. Note that RanGAP expression values shown are log2 >9.0. Heat maps were constructed using BAR HeatMapper Plus tool (http://bar.utoronto.ca). See Zimmermann et al., 2004 for anatomical definitions.

107 A RanGAP1::GUS B RanGAP1::GUS

C RanGAP1::GUS D RanGAP2::GUS

Figure 2.13: Assay for RanGAP promoter-GUS expression shows a high level of staining in the vegetative meristems of shoot and roots. (A, B): GUS staining for the activity of RanGAP1 promoter in (A) primary shoot and (B) primary root of transgenic 3 dpg seedlings. Arrow in (A) points to the intense staining in the SAM and arrow in (B) points to the intense staining in the RAM. (C, D): GUS staining for the activity of RanGAP promoters in 6 dpg seedlings. (C) Arrow points to intense GUS staining in the lateral root initiating cells (i.e. meristematic cells in the pericycle) at the base of a developing lateral root primordium in a RanGAP1 promoter-GUS seedling. (D) shows a lateral root with prominent staining in the lateral root meristem in a RanGAP2 promoter- GUS seedling. Note that RanGAP promoter activity is not restricted to the meristem regions. Data generated by Wang, H-J. Scale bars: 100 µm. Seedlings were grown on 2% sucrose (w/v) MS medium.

108

Figure 2.14: Phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 are rescued by native promoter- driven genomic constructs. (A) Phenotypic recovery of a 12 dpg rg1-1/rg1-1;rg2-2/rg2- 3 seedling carrying the native RanGAP1 promoter-driven genomic construct compared to rg1-1/rg1-1;rg2-2/rg2-3 and wild-type (Col). (B) Immunoblot analysis of phenotypically recovered rg1-1/rg1-1;rg2-2/rg2-3 showing expression of RanGAP1 from the genomic construct. Total protein extracts from 12 day-old seedlings were incubated with the anti- RanGAP1 antibody. Coomassie brilliant blue staining of a replica gel is shown at the bottom as the loading control. Line numbers refer to the transgenic rg1-1/rg1-1;rg2- 2/rg2-2 lines carrying the native promoter construct that were used for the crosses. Seedlings were grown on 2% sucrose (w/v) MS medium.

109 Figure 2.14

rg1-1/rg1-1;

rg1-1/rg1-1; rg2-2/rg2-3 + RG1 Col WT rg2-2/rg2-3 Line 1

rg1-1/rg1-1; rg1-1/rg1-1; rg2-2/rg2-3 + RG1 rg2-2/rg2-3 Line 1 Line 2 Col WT

α-RG1

Coomassie

110

Figure 2.15: Nuclear import is not impeded in rg1-1/rg1-1;rg2-2/rg2-3. Confocal microscopy images showing the localization of GFP-N7 nuclear import marker (Cutler et al., 2000; Cutler and Somerville, 2005) in the transgenic (A, C) rg1-1/rg1-1;rg2-2/rg2-3 and (B, D) rg1-1/rg1-1;RG2/rg2-2 seedlings at 11 dpg. (C) and (D) were captured at greater resolution for closer observation. GFP-N7 is nuclear localized and cytoplasmic accumulation of GFP-N7 is not seen in (A, C) rg1-1/rg1-1;rg2-2/rg2-3 similar to (B, D) rg1-1/rg1-1;RG2/rg2-2. (E) serves as a control. In (E), nuclear import of the GFP-N7 marker is evaluated in the inducible GDP-locked mutant (Ran1T27N) and GTP-locked mutant (Ran1Q72L) that disrupt the Ran cycle. GFP florescence (i.e. GFP-N7 accumulation) is seen in the cytoplasm under induced condition indicating perturbed nuclear import of the GFP-N7 marker. Details of the mutant constructs and transgenic plants are described in Materials and methods. (F)-(I) show nuclear localization of the nuclear import marker GFP-WIP1ΔTDF (Xu et al., 2007b) in root tip cells of an induced RanGAP RNAi line (RanGAP2RNAi/rg1-1) and cytoplasmic localization of GFP- WIP1ΔTDF fluorescence is not observed. (H) contains morphologically unaltered cells. (I) shows a morphologically disrupted cell [i.e. a swollen cell]. Images (H) and (I) show propidium-iodide stained cell walls (magenta color) of the images presented in (F) and (G) respectively. (E)-(I) were generated by Xu, X.M. Scale bars in A and C: 50 µm, B, D, E-I: 20 µm. Seedlings were grown on 2% sucrose (w/v) MS medium.

111 Figure 2.15

rg1-1/rg1-1; rg1-1/rg1-1; rg2-2/rg2-3 RG2/rg2-2 A B RanGAP2RNAi/rg1-1 F G

C D HI

112 A B

rg1-1/rg1-1;rg2-2/rg2-3 rg1-1/rg1-1;RG2/rg2-2

C D

rg1-1/rg1-1; rg1-1/rg1-1; rg2-2/rg2-3 RG2/rg2-2

Figure 2.16: CycB1;1::GUS expression is restricted to smaller regions in the shoot and root apices of rg1-1/rg1-1;rg2-2/rg2-3 compared to rg1-1/rg1-1;RG2/rg2-2. GUS assay of transgenic 5 dpg seedlings containing the mitotic marker CycB1;1::GUS construct. CycB1;1::GUS expression indicates regions with cell division activity. GUS staining is restricted to a smaller area in (A and C) rg1-1/rg1-1;rg2-2/rg2-3 compared to (B and D) rg1-1/rg1-1;RG2/rg2-2. (A and B) shoot apices; (C and D) root apices. Comparative pairs; (A)/(B) and (C)/(D) are presented at the same magnification. Scale bars: 100 µm. Seedlings were grown on 2% sucrose (w/v) MS medium.

113

Figure 2.17: Growth and developmental phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 seedlings are rescued by sucrose supplementation. rg1-1/rg1-1;rg2-2/rg2-3 and rg1-1/rg1-1;RG2/rg2-2 seedlings grown in MS medium (A) lacking sucrose and (B) containing 3% sucrose (w/v) at 12 dpg. rg1-1/rg1-1;rg2-2/rg2-3 seedlings that failed to establish in sucrose-lacking medium are shown in (A). In comparison, rg1-1/rg1- 1;rg2-2/rg2-3 achieved near wild-type recovery in shoot and root phenotypes with 3% sucrose supplementation as seen in (B). Note that rg1-1/rg1-1;RG2/rg2-2 seedlings lack significant growth differences in the media lacking sucrose and containing 3% sucrose at 12 dpg. A representative genotyping result of rg1-1/rg1- 1;rg2-2/rg2-3 and rg1-1/rg1-1; RG2/rg2-2 on 0% sucrose and 3% sucrose are shown in bottom panels of (A) and (B). Arrows indicate the expected position for the PCR product of the tested allele. For primer details see Materials and methods.

114 Figure 2.17

0% Sucrose 3% Sucrose A B

rg1-1/rg1-1; rg1-1/rg1-1; rg1-1/rg1-1; rg1-1/rg1-1; RG2/rg2-2 rg2-2/rg2-3 RG2/rg2-2 rg2-2/rg2-3

rg1-1/rg1-1; rg1-1/rg1-1; rg1-1/rg1-1; RG2/rg2-2 rg1-1/rg1-1;rg2-2/rg2-3 RG2/rg2-2 rg2-2/rg2-3

RG2 rg2-3 RG2 rg2-3 RG2 rg2-3 RG2 rg2-3

Plant 1 Plant 2 Plant 1 Plant 2

115

Figure 2.18: Adult shoot phenotypes of rg1-1/rg1-1;rg2-2/rg2-3 are rescued by sucrose supplementation. (A)-Top panel: rg1-1/rg1-1;RG2/rg2-2 (left) and rg1-1/rg1-1; rg2-2/rg2-3 (right) seedlings at 21 dpg grown in solid MS medium containing 3% sucrose (w/v) for 14 days followed by transfer to liquid MS medium containing 3% sucrose (w/v). Radial leaf arrangement is present in both rg1-1/rg1-1;RG2/rg2-2 and rg1- 1/rg1-1;rg2-2/rg2-3, and the rosette sizes do not deviate considerably. In comparison, a 21 dpg rg1-1/rg1-1;rg2-2/rg2-3 seedling grown on 1.5% sucrose (w/v) MS medium [(A)- Bottom panel] contains a more compact shoot consisting of distorted phyllotaxy and multiple small rosettes. Arrow points to a mal-formed and crinkled first true leaf. Scale bars: 17 mm. (B) rg1-1/rg1-1;rg2-2/rg2-3 grown on 3% sucrose (w/v) MS medium up to 14 dpg and transferred to soil (middle) achieves the height and stature of a similarly grown rg1-1/rg1-1;RG2/rg2-2. In comparison, a rg1-1/rg1-1;rg2-2/rg2-3 grown on 1.5% sucrose (w/v) MS medium up to 14 dpg and transferred to soil (right) shows shorter height and bushy phenotype. Plants were photographed ~55 dpg. Siliques of rg1-1/rg1- 1;rg2-2/rg2-3 transferred from the 3% sucrose medium are small and similar in size to siliques of rg1-1/rg1-1;rg2-2/rg2-3 transferred from the 1.5% sucrose medium (i.e. siliques are not complemented by sucrose supplementation). (C) Enlarged view showing the rosettes of plants in (B). The rosette of rg1-1/rg1-1;rg2-2/rg2-3 transferred from 3% sucrose medium achieves the size and ordered leaf arrangement similar to rg1-1/rg1- 1;RG2/rg2-2 but the rg1-1/rg1-1;rg2-2/rg2-3 transferred from 1.5% sucrose medium consists of a smaller shoot made of multiple compact adventitious rosettes. (D) Enlarged view of plants in (B) showing the smaller siliques of rg1-1/rg1-1;rg2-2/rg2-3 compared to rg1-1/rg1-1;RG2/rg2-2.

116 Figure 2.18

A B 3% sucrose

rg1-1/rg1-1; rg1-1/rg1-1; RG2/rg2-2 rg2-2/rg2-3

1.5% sucrose

rg1-1/rg1-1; rg2-2/rg2-3

3% sucrose 3% sucrose 1.5% sucrose rg1-1/rg1-1; rg1-1/rg1-1;rg2-2/rg2-3 RG2/rg2-2 Continued

117 Figure 2.18: Continued

3% sucrose 3% sucrose 1.5% sucrose rg1-1/rg1-1; rg1-1/rg1-1;rg2-2/rg2-3 RG2/rg2-2

D 3% sucrose

rg1-1/rg1-1; RG2/rg2-2

rg1-1/rg1-1; rg2-2/rg2-3

118

Figure 2.19: Cell numbers at the root tip and root morphology are rescued by sucrose supplementation. FM4-64 (8 dpg, A-D) and propidium iodide (6 dpg, E-H) stained root tips were optically sectioned by confocal microscopy to reveal histology at a sub-surface plane (A-D) or the mid plane (E-H). Seedlings were grown in MS medium lacking sucrose (A, C, E, G) or containing 3% sucrose (w/v) (B, D, F, H). rg1-1/rg1- 1;rg2-2/rg2-3 seedlings are shown in panels (A), (B), (E) and (F). rg1-1/rg1-1;rg2-2/rg2- 3 in panels (A) and (E) are halted in growth prior to seedling establishment. rg1-1/rg1- 1;RG2/rg2-2 seedlings are shown in (C), (D), (G) and (H). Dotted lines in (A) and (E) indicate the boundary between the meristematic and cell elongation zones. In all other panels this boundary lies outside of the pictured area. Differences in the overall root tip morphology of rg1-1/rg1-1;rg2-2/rg2-3 (A and E) compared to rg1-1/rg1-1;RG2/rg2-2 (C and G) on 0% sucrose medium are seen owing to the small size of root, reduced number of cells, compact cell division zone and placement of longitudinally elongating cells closer to the root tip in rg1-1/rg1-1;rg2-2/rg2-3. With 3% sucrose supplementation, the cell numbers and overall root tip morphology are restored in rg1-1/rg1-1;rg2-2/rg2-3 (B, F) to resemble rg1-1/rg1-1;RG2/rg2-2 (D, H). Note that the cell numbers and overall root tip morphology are indistinguishable between rg1-1/rg1-1;RG2/rg2-2 seedlings grown on 0% sucrose (C and G) and 3% sucrose media (D and H). Arrow in (E) points to new root cap cells of rg1-1/rg1-1;rg2-2/rg2-3 on 0% sucrose medium which are misaligned with the older root cap cells and are of irregular shapes (i.e indicates beginning of aberrant cell division towards the root cap). Oblique cell walls, irregular cell shapes and disorganized cell files in the root cap region of rg1-1/rg1-1;rg2-2/rg2-3 are seen to persist upon sucrose supplementation (examples shown by arrowheads in B and F). All panels are presented at the same magnification. Scale bars: 100 µm.

119 Figure 2.19: Continued

0% sucrose 3% sucrose A B

rg1-1/rg1-1; rg2-2/rg2-3

> >

C D

rg1-1/rg1-1; RG2/rg2-2

Sub-surface plane

Continued

120 Figure 2.19: Continued

0% sucrose 3% sucrose E F

rg1-1/rg1-1; rg2-2/rg2-3

H G

rg1-1/rg1-1; RG2/rg2-2

Mid plane

121

Figure 2.20: Sucrose supplementation rescues CycB1;1::GUS expression in the shoot apex of rg1-1/rg1-1;rg2-2/rg2-3. GUS staining of 3 dpg seedlings of transgenic (A) rg1- 1/rg1-1;rg2-2/rg2-3 and (B) rg1-1/rg1-1;Rg2/rg2-2 containing the mitotic marker CycB1;1::GUS and grown on MS medium lacking sucrose. CycB1;1::GUS expression indicates regions with cell division activity. (C) shows representative rg1-1/rg1-1;rg2- 2/rg2-3 or rg1-1/rg1-1;Rg2/rg2-2 seedlings grown on MS medium containing 3% sucrose (w/v). Among >60 seedlings stained, no seedling was observed that would resemble the seedling shown in (A). Note that first true leaves are not observable in (A) but are developed in (B) and (C). GUS staining shows a similar pattern (i.e. meristem and young leaves) in the shoot apices of (B) and (C). Robustness of the seedling in (B) is due to developmental variation present among seedlings of rg1-1/rg1-1;Rg2/rg2-2 which is independent of sucrose concentration. All four panels are presented at the same magnification. Scale bars: 100 µm.

122 Figure 2.20

0% sucrose

rg1-1/rg1-1;rg2-2/rg2-3 rg1-1/rg1-1;RG2/rg2-2

A B

3% sucrose C

123 0% sucrose 3% sucrose

A B

rg1-1/rg1-1; rg2-2/rg2-3

C D

rg1-1/rg1-1; RG2/rg2-2

Figure 2.21: Sucrose supplementation rescues CycB1;1::GUS expression in the root apex of rg1-1/rg1-1;rg2-2/rg2-3. GUS staining of 6 dpg seedlings of transgenic (A, B) rg1-1/rg1-1;rg2-2/rg2-3 and (C, D) rg1-1/rg1-1;Rg2/rg2-2 containing the mitotic marker CycB1;1::GUS construct. CycB1;1::GUS expression indicates regions with cell division activity. Seedlings were grown in MS medium (A, C) lacking sucrose or (B, D) containing 3% sucrose (w/v). Note that the root size is smaller in (A). Roots in (B)-(D) show similar size and morphology. GUS staining is barely observable in (A). GUS staining is similar among root apices (B)-(D). All four panels are presented at the same magnification. Scale bars: 200 µm.

124 Table 2.1: Silique data from rg1-1/rg1-1;RG2/rg2-2 and rg1-1/rg1-1;rg2-2/rg2-3 plants

Parent: rg1-1/rg1-1;RG2/rg2-2 Parent: rg1-1/rg1-1;rg2-2/rg2-3

Developed Aborted Aborted Total Silique Developed Aborted Aborted Total Silique seeds ovules seeds length seeds ovules seeds length (cm) (cm) 54 0 1 55 1.35 0 23 0 23 0.20

56 1 0 57 1.40 0 22 1 23 0.25

51 2 0 53 1.40 0 22 0 22 0.25

51 3 0 54 1.30 0 24 0 24 0.20

50 0 0 50 1.40 0 23 0 23 0.25

‘Total’ refers to the sum of developed seeds, aborted ovules and aborted seeds (i.e. total number of ovule primordia initiated).

125 Table 2.2: Transmission of the rg1-1;rg2-3 genotype in comparison to rg1-1;rg2-2 through the male gametophyte

Cross RG1/rg1-1; RG1/rg1-1; Number of TE RG2/rg2-2 RG2/rg2-3 progeny Male Female Male genotyped %

RG1/RG1; rg1-1/rg1-1; 138 107 245 77.5 RG2/RG2 rg2-2/rg2-3

TE, transmission efficiency. TE was calculated according to Howden et al., 1998. TE= progeny containing rg1-1;rg2-3 /progeny lacking rg1-1;rg2-3 X 100%.

126 CHAPTER 3

INVESTIGATION OF PROTEIN-PROTEIN INTERACTIONS AND STRESS RESPONSE–RELATED PHOSPHORYLATION OF TANDEM ZINC FINGER 1 (TZF1) IN ARABIDOPSIS THALIANA

127 3.1 Abstract

Tandem Zinc Finger 1 of Arabidopsis thaliana (AtTZF1) is a plant-unique CCCH-type tandem zinc finger (TZF) protein implicated in drought and cold stress tolerance via regulation of gene expression. Stress-responsive cytoplasmic foci consisting of mRNA and proteins; stress granules (SGs) and processing bodies (PBs) are implicated in AtTZF1-mediated regulation of gene expression. Regulation of AtTZF1 subcellular localization and the dynamics of SG and PB assembly are postulated to be important for AtTZF1 functions, although the exact molecular mechanisms remain elusive. Microarray- based experiments have shown that AtTZF1 is a positive regulator of ABA-mediated gene expression which is consistent with its stress tolerance functions. Studies with the most well studied CCCH-type tandem zinc finger protein in human; hTTP, has revealed regulation of hTTP activity via the Mitogen-Activaed Protein Kinase (MAPK) pathway in stress response. Further, stress response studies with the cotton TZF; GhZFP1 has revealed direct interacting partners that may be involved in mediating GhZFP1 stress- combat functions.

In an effort to understand the molecular mechanisms underlying the functions of AtTZF1 including stress response, candidate interactions for AtTZF1 based on a yeast two-hybrid (Y-2-H) screen and co-localization experiments were pursued in this study. All tested putative interacting partners failed to establish a direct interaction using the full length clones in the vector system employed. Because sequence analysis revealed that AtTZF1 contains a MAPK docking site, and the possibility that MAPK pathway may be involved in regulating AtTZF1 in stress response, one of the candidates; AtMPK11 (Arabidopsis thaliana Mitogen-activaed Protein Kinase 11) was further tested for its ability to phosphorylate AtTZF1 via an in vitro phosphorylation assay. The results revealed that AtMPK11 does not phosphorylate AtTZF1 under the conditions tested. Alternative and/or improved assays are necessary to further evaluate these results.

128 3.2 Introduction

3.2.1 Zinc fingers, diversity and importance

An overview of Zinc (Zn) finger proteins is presented in Iuchi and Kuldell, 2005. A Zn finger is a peptide domain with a special tertiary structure stabilized by coordination of cysteine (C) and histidine (H) residues to a Zn (II) ion (Vargek et al., 1999). The first discovered and most abundant are the C2H2-type Zn fingers which are present in ~2% (~700 genes) of human genome and referred to as the classical Zn finger proteins. Many of the C2H2-type Zn finger proteins are transcription factors. Some C2H2 Zn fingers are also known to mediate protein-protein interactions to result in selective homo- and hetero-dimerization (McCarty et al., 2003). Later, many other Zn finger proteins with variants in the Zn finger motif were identified such as C2C2, CCHC, CCCH, LIM, RING, TAZ and FYVE which are classified in different subclasses. These Zn fingers may occur singly or as tandem repeats of varying numbers.

The tertiary structures of the Zn fingers confer specific binding properties to various molecules such as DNA, RNA, proteins and small molecules, and the functions of Zn finger proteins include gene expression, signal transduction, cell growth, differentiation and development. Defects in some of these genes are found associated with disease such as cancer and neurological diseases. Zn deficiency leads to cell death probably due to the importance of Zn finger proteins in cellular functions. The nucleic acid binding property of Zn fingers is exploited by scientists to specifically target Zn finger molecules to DNA or RNA of interest which is an area of research in developing designer drugs to control the expression of disease associated genes (Laity et al., 2001).

129 3.2.2 CCCH Zn finger proteins and CCCH-type Tandem Zn Finger proteins

CCCH Zn fingers are abundantly represented in the eukaryote genomes from yeast to animals and plants including budding yeast, fission yeast, human, mouse, rice, soybean and Arabidopsis (Cao, 2008; Pomeranz et al., 2011a). CCCH Zn finger domain is diverse in the number of tandem Zn fingers present in the protein ranging from one to six, as well as in the structure of the Zn fingers itself (Liang et al., 2008). Tandem Zn Finger (TZF) proteins are characterized by two tandem CCCH-type Zn fingers. Human TZF family consists of 3 genes; Zfp36, Zfp36l1 and Zfp36l2 and the most well studied member; Zfp36 is more commonly known as tristetraproline (TTP) (Carrick et al., 2004; Lai et al., 1990).

CCCH-type TZFs are involved in binding to nucleic acid (DNA and RNA), protein and small molecules (Iuchi and Kuldell, 2005; Pomeranz et al., 2010a). It is postulated that TZF proteins are likely mediators of cell signaling where their effects are brought on via these molecular interactions. They play important roles in gene expression, cell fate determination, developmental and environmental responses (Mello et al., 1996; Blackshear et al., 2005; Lin et al., 2011; Pomeranz et al., 2011a). The three human TZF proteins are nuclear-cytoplasmic shuttling proteins (Phillips et al., 2002).

3.2.3 Cellular and molecular mechanisms mediating TZF functions

Involvement of TTP as a signaling component in growth and stress response that brings about regulation of gene expression is reviewed by Dean et al., 2004 and Cao, 2008. This includes I) transcriptional and post-translational regulation of TTP via upstream signaling events, and II) downstream processes regulated by TTP.

The TTP mRNA levels and TTP protein stability are increased in response to growth and stress stimuli. These include insulin and other growth factors as well as stimulators of

130 innate immunity such as the endotoxin and bacterial lipopolysaccharide. Further, TTP is a hyperphosphorylated protein (Cao et al., 2006) and phosphorylation has been shown to act as a molecular switch for TTP activity. For instance, stress-induced kinase signaling cascades, particularly Mitogen-Activated Protein Kinase (MAPK) pathway is known to affect the phosphorylation status of TTP (Dean et al., 2004; Stoecklin et al., 2004, Taylor et al., 1995), thereby determining its downstream molecular functions.

Subcellular localization of TTP is important for TTP functions. Rapid translocation of TTP from nucleus to cytoplasm results in response to stimuli such as mitogens (Taylor et al., 1996). Further, phosphorylation of TTP promotes its nuclear export (Johnson et al., 2002), thereby facilitating its functions in the cytoplasm. TTP localizes to cytoplasmic foci known as p-bodies (PBs) and under stress conditions to stress granules (SGs) (Kedersha et al., 2005). PBs and SGs are dynamic cytoplasmic foci with discrete and overlapping functions (Anderson and Kedersha, 2008). PBs and SGs are assembled in stress response by TTP and other RNA-binding proteins that contain protein-protein interaction domains (Kedersha et al., 2005; Franks and Lykke-Andersen, 2007; Anderson and Kedersha, 2008). They are the sites for TTP-mediated translational repression (Franks and Lykke-Andersen, 2007) and mRNA decay (Anderson and Kedersha, 2008; Anderson and Kedersha, 2009) and are present in eukaryotes from yeast (Teixeira and Parker, 2007; Buchan et al., 2008) to animals (Anderson and Kedersha, 2002; Sheth and Parker, 2003; Cougot et al., 2004) to plants (Pomeranz et al., 2010a). PBs and SGs are structurally, functionally and spatially linked (Kedersha et al., 2005; Anderson and Kedersha, 2008). However, P-bodies are primarily involved in mRNA degradation while stress granules are sites of mRNA sorting that contain stalled translational pre-initiation complexes (Valencia-Sanchez et al., 2006). Triage in stress granules could direct mRNA in the stalled pre-initiation complexes to resume translation or undergo degradation in PBs depending on the stress-combat situation (Valencia-Sanchez et al., 2006). The fate of each mRNA (i.e. whether recruited to SGs or preferentially associated with polysomes, destiny upon triage in SGs such as degradation, retention in a translationally repressed

131 state or resume translation) is a selective process depending on multiple factors including the functions of the encoded proteins and cellular needs for their functions at a given time (Anderson and Kedersha, 2008).

With regard to the molecular mechanisms by which TTP executes its functions, TTP silences target mRNAs by mRNA decay and translational repression, both of which are mechanisms of post-transcriptional gene regulation (Franks and Lykke-Andersen, 2007). Human TTP (hTTP) binds to type II AU rich elements (AREs) at the 3’UTR of target mRNA and interact via its N-terminal activation domain with enzymes involved in deadenylation, decapping and exonucleolytic activities (Carballo et al., 1998; Lai et al., 1999; Carrick et al., 2004; Lykke-Andersen and Wagner, 2005; Liang et al., 2008). Thus these mRNA decay machineries are recruited to target ARE-containing mRNA by TTP. When binding to class II AU rich elements of mRNA, the two Zn fingers are known to bind the two identical halves of the ARE region (Carrick et al., 2004; Hudson et al., 2004; Barreau et al., 2006). While deadenylases remove the poly(A) tail of the target mRNA, the decapping enzyme complex removes the 5’ cap. Subsequently, 5’ to 3’ mRNA degradation takes place via a specific exonuclease, XRN1 (Wilusz and Wilusz, 2004; Lykke-Andersen and Wagner, 2005; Anderson and Kedersha, 2006). A similar mechanism of ARE-mediated mRNA degradation is postulated to exist in other animals, plants and yeast as well (Wilusz and Wilusz, 2004; Anderson and Kedersha, 2006; Pomeranz et al., 2010a)

Of the human genome, 5-8% of the genes contain AREs (Bakheet et al., 2003; Wilusz and Wilusz, 2004) that could be potential targets for the TTP-mediated mRNA turnover. In animals, the deficiency of TTP lead to a number of inflammatory disorders including arthritis, autoimmunity and cardiovascular diseases likely due to excessive production of pro-inflammatory cytokines TNFα (Tumor necrosis factor-α; a cytokine involved in inflammation) and GM-CSF (granulocyte–macrophage colony stimulating factor that

132 contains a TNFα-like ARE), whose mRNA are direct targets of TTP (Carrick et al., 2004; Cao, 2008).

MAPK pathway converts extra-cellular stimuli to specific cellular responses via a phosphorelay cascade and is involved in communicating stress signals in eukaryotes from fungi to animals to plants (Colcombet and Hirt, 2008). Activation of the stress-induced p38-MAPK and c-Jun N terminal kinase (JNK) pathways has shown to stabilize ARE- containing mRNAs (Ming et al., 1998; Winzen et al., 1999). During arsenite-induced oxidative stress in mammalian cell cultures, phosphorylation of hTTP at Ser52 and Ser178 by the p38-MAPK pathway promotes binding of 14-3-3 (Stoecklin et al., 2004). The cytoplasmic interaction of hTTP and 14-3-3 prevents the localization of hTTP to SGs and thereby inhibits degradation of ARE-containing mRNA (Stoecklin et al., 2004). 14-3- 3, a chaperon protein which stabilizes proteins in certain conformations, interacts with phospho-serines of selective partners and thereby elicits a wide range of cellular effects (Tzivion and Avruch, 2002; Wilusz and Wilusz, 2004). In hTTP, 14-3-3 binds to phophorylated Ser 178 (Johnson et al., 2002). However, in mitochondrial stress induced by a mitochondrial inhibitor, the p38-MAPK and JNK pathways are not activated (Stoecklin et al., 2004), hTTP remains unphosphorylated, localizes to SGs and degrades ARE-containing mRNA (Wilusz and Wilusz, 2004).

Emerging data suggest that TTP may be involved in microRNA-mediated mRNA decay that takes place in the PBs (Jing et al., 2005). A human microRNA; miR16 has shown to require hTTP to perform microRNA-mediated decay of TNF-α mRNA in HeLa cells, yet does not directly interact with hTTP. However, co-immunoprecipitation experiments have shown association of hTTP with miR16 and the endonuclease AGO2 of the RNA- induced silencing complex (RISC) in the RNA interference machinery. Therefore, it is postulated that TTP may facilitate microRNA mediated mRNA decay via interaction with the components of the RISC (Jing et al., 2005). Further experiments suggest that this mode of mRNA decay may be different from the usual slicer-based microRNA-mediated

133 mRNA decay pathway (Valencia-Sanchez et al., 2006). In an independent study, TTP and its paralog Zfp36l1/BRF-1 have shown to decay a non-ARE containing reporter mRNA (Lykke-Andersen and Wagner, 2005) of which the mechanism remains to be studied. Further, hTTP has been implicated in exosome-mediated 3’ to 5’ decay of mRNA via the recruitment of exosome subunits to the ARE-containing mRNA (Chen et al., 2001).

The mechanism of translational repression via TTP is not well understood. Sequestering of target ARE containing mRNAs in PBs and SGs by TTP maintains the recruited mRNAs in a polysome-free translationally repressed state. This process is enhanced when mRNA decay enzymes are limiting in the cytoplasm suggesting that TTP functions to translationally silence mRNA by associating them in cytoplamic bodies (Franks and Lykke-Andersen, 2007). Translational regulatory proteins are localized in PBs and SGs including the translational repressors 4E-T, p54/RCK, TIAR and TIA-1 (Andrei et al., 2005; Ferraiuolo et al., 2005; Anderson and Kedersha, 2006; Kedersha et al., 1999; Kedersha et al., 2005; Mazan-Mamczarz et al., 2006). However, the underlying molecular mechanisms may be coupled with other mRNA silencing pathways such as mRNA deadenylation and microRNA machinery (Franks and Lykke-Andersen, 2007). Endonuclease-mediated mRNA cleavage and/or mRNA decay-independent translational repression has been proposed for microRNA-based gene regulation (Valencia-Sanchez et al., 2006).

Alternative molecular mechanisms have been indicated for other CCCH-type Zn finger proteins in other organisms. For example, a unique CCCH Zn finger protein in rice (OsLIC) has been indicated in transcriptional activation activity (Wang et al., 2008b). PIE-1 in Caenorhabditis elegans, a protein containing a TZF motif and expressed in germline blastomeres have been indicated in repression of transcription (Mello et al., 1996). Stress tolerance conferred by the cotton TZF; GhZFP1 has been indicated to occur via interaction with stress responsive proteins; RD21A and PR5 (Guo et al., 2009). In the stress responsive activities by AtSZF1 (AtTZF11) / AtSZF2 (AtTZF10), a known

134 mechanism for TZF-mediated mRNA decay in PBs (i.e. via ARE element or microRNA) can not be assumed because AtSZF1 has been shown to localize to the nucleus but not to PBs (Sun et al., 2007). Nevertheless, Pomeranz et al., 2010b showed that AtTZF10 and AtTZF11 localized in cytoplasmic foci in a maize protoplast transient expression system.

3.2.4 Arabidopsis CCCH-type Tandem Zn Finger (TZF) proteins

CCCH-type Zn finger family is described in Blackshear et al., 2005; Liang et al., 2008; Wang et al., 2008a; Pomeranz et al., 2010a. Sixty eight CCCH Zn finger genes have been identified in a genome-wide screen in Arabidopsis thaliana which are grouped into 11 subfamilies. Among them, 26 genes contain a TZF motif. However, only two contain the

same TZF motifs as in hTTP which is C-X8-C-X5-C-X3-H-X18-C-X8-C-X5-C-X3-H (i.e.

two identical C-X8-C-X5-C-X3-H motifs separated by 18 amino acids, X denotes any amino acid). By contrast, 11 out of these 26 genes contain a unique TZF motif that can only be found in plants and constitute the subfamily IX. This plant unique motif contains

a C-X5-C-X4-C-X3-H configuration in the second CCCH Zn finger and is preceded by a

first C-X7-8-C-X5-C-X3-H Zn finger separated by 16 amino acids. Additionally, a ~50 amino acid region upstream of the TZF motif remains plant unique and highly conserved among the members of the Arabidospis subfamily IX genes. Similar TZF motif structure and upstream plant-unique region are conserved in the majority of genes in rice TZF subfamily I as well. The plant-unique region contains a putative CHCH Zn finger motif

(C-X5-H-X4-C-X3-H) spaced invariably in these genes (i.e. 19 amino acids upstream of the first CCCH Zn finger), which may imply an associated biological function (For details see Blackshear et al., 2005; Liang et al., 2008; Wang et al., 2008a; Pomeranz et al., 2010a).

Proteins encoded by the 11 genes of Arabidopsis thaliana TZF subfamily IX are designated AtTZF1-11 (Pomeranz et al., 2010a). Functions of only a few of these proteins are known. PEI1 (AtTZF6) has shown to be important for embryo development

135 (Li and Thomas, 1998) while AtSZF1 (AtTZF11) and AtSZF2 (AtTZF10) enhance salt stress tolerance (Sun et al., 2007). SOMNUS (AtTZF4) is associated with light-dependent seed germination (Kim et al., 2008).

3.2.5 AtTZF1 structure, functional implications and stress response

AtTZF1 (AtCTH/AtC3H23) focused in this study was identified in a screen for sugar responsive genes (Price et al., 2004). The AtTZF1 gene; At2g25900 encodes a protein of 315 amino acids with a predicted molecular weight of ~35.46 kD [The Arabidopsis Information Resource (TAIR, http://www.arabidopsis.org/)]. It contains the configuration of C-X7-C-X5-C-X3-H-X16-C-X5-C-X4-C-X3-H in the TZF motif and the upstream

putative CHCH Zn finger (C-X5-H-X4-C-X3-H), all of which are plant unique as described above (Liang et al., 2008; Wang et al., 2008a; Pomeranz et al., 2010a). Localization experiments with GFP-fusion AtTZF1 protein in plant protoplasts showed that it shuttles between the nucleus and cytoplasm, and localizes to cytoplasmic foci similar to hTTP (Pomeranz et al., 2010a; Pomeranz et al., 2010b). AtTZF1-localizing cytoplasmic foci are also present in specific tissues of intact plants and are stress- inducible (Pomeranz et al., 2010a). Studies in plant protoplasts have shown that in these cytoplasmic foci, AtTZF1 co-localize with PB markers; DCP2 (At5g13570), XRN4 (At1g54490) and AGO1 (At1g48410), and under heat-induced stress conditions with the SG marker; PABP8 (At1g49760). These results indicate a PB- and SG- identity for these cytoplasmic foci and suggest that AtTZF1 may share similar roles with TTP in PBs and SGs. Consistent with the known mechanisms of function for TZF proteins described above that requires binding to nucleic acids (i.e. mRNA decay, transcriptional regulation and translational repression), in vitro binding assays demonstrated that AtTZF1 binds both DNA and RNA in a Zn-dependent manner (Pomeranz et al., 2010a). However, preliminary assays in yeast and maize protoplasts suggest that AtTZF1 might not function as a transcriptional activator or repressor (Pomeranz et al., 2011b). It was also shown that AtTZF1 does not bind to hTTP-specific 5’-UAUUUAUU-3’ consistent with the idea that

136 specificity of binding to an ARE is dependant upon the conformation of the Zn fingers (Pomeranz et al., 2010a).

In general the subfamily IX TZF genes are responsive to stress conditions including osmotic and cold conditions (Liang et al., 2008; Wang et al., 2008a; Pomeranz et al., 2010a; Lin et al., 2011). Upregulation of salt responsive genes and increased sensitivity to salt stress were shown by mutants of AtTZF10 and AtTZF11, while the opposite trends were shown by AtTZF11-overexpression plants (Sun et al., 2007). However, the underlying molecular mechanisms with respect to AtTZF10 and AtTZF11 remain to be elucidated. On the other hand, tobacco plants overexpressing cotton GhZFP1, a TZF closest to AtTZF4 and AtTZF5 showed abiotic (salinity) and biotic (fungal infestation) stress tolerance (Guo et al., 2009). Although the details of the molecular mechanism are unknown, a yeast two-hybrid (Y-2-H) screen has reveled interaction of GhZFP1 with RD21A (responsive to dehydration protein 21A, a cysteine-type proteinase responsive to water deprivation) and PR5 (pathogenesis-related protein 5, a component of the plant defense system) (Guo et al., 2009) which are proteins involved in stress response.

Loss-of-function studies of AtTZF1 revealed that T-DNA knockout plants lack noticeable vegetative plant-phenotypes (Lin et al., 2011). However, AtTZF1-overexpression (OE) plants show compact rosette phenotypes during the first few weeks post-germination. This defect recovers from ~6-8 weeks of age and subsequently surpasses the biomass of the wild-type plants. AtTZF1-OE plants are late flowering and longevity of the plants is increased (Lin et al., 2011). Interestingly, AtTZF1-OE plants show drought- and freeze stress tolerance in non-acclimated plants, while RNAi plants with loss of function of AtTZF1, AtTZF2 and AtTZF3 show higher water loss (Lin et al., 2011).

ABA plays a critical role in a variety of stress-signaling including drought and cold (Xiong et al., 2002). ABA and GA are antagonistic for most physiological effects. Molecular evidence reveals that ABA stabilizes DELLA proteins which are ‘GA effect’

137 repressors (Tyler et al., 2004; Pareek et al., 2010). The stress-tolerance phenotypes of AtTZF1-OE plants are suggestive of ABA over-accumulation and GA deficiency. However, the ABA and GA levels were not significantly altered in the AtTZF1-OE plants under drought or freezing conditions (Lin et al., 2011), which indicates that AtTZF1 may not influence the metabolism of these hormones unlike SOMNUS (AtTZF4) (Kim et al., 2008). In loss-of-function mutants of SOMNUS, the ABA level was reduced and the GA level was elevated via expression changes in the ABA and GA metabolic genes (Kim et al., 2008). On the contrary, the RNAi mutant of AtTZF1-3 also showed unchanged ABA and GA levels compared to wild-type and AtTZF1-OE plants (Lin et al., 2011). Yet, microarray based expression analysis have shown that ABA responsive genes are upregulated while the GA responsive genes are downregulated in the AtTZF1-OE plants (Lin et al., 2011). These results together suggest that AtTZF1 may act as a downstream molecular switch that positively regulate ABA responses and negatively regulate GA responses via gene expression (Lin et al., 2011). However, the exact mechanisms of action and the direct downstream targets of AtTZF1 are currently unknown. Investigating the functions of TZF1 in planta is particularly interesting due to its involvement in the phytohormonal signaling pathways (i.e. possibly acting as a molecular switch downstream of ABA/GA) and environmental stress conditions (i.e. cold and drought tolerance of AtTZF1-OE plants) which may unravel how its gene-regulation mechanism(s) are coupled with the plant-unique signaling network.

Several indirect evidences suggest possible regulation of AtTZF1 by the MAPK pathway. Firstly, TTP is regulated by p38-MAPK pathway-mediated phosphorylation in human stress response (see section 3.2.3). Secondly, both the plant MAPK pathway and AtTZF1-OE plants are implicated in stress response, including cold and drought tolerance (Colcombet and Hirt, 2008; Pomeranz et al., 2010a). Supporting this hypothesis in a more direct manner, a report based on a protein microarray study that screened for substrates of ten AtMPKs (i.e. AtMPKs 1-8, AtMPK10 and AtMPK16) indicated AtTZF1 as a substrate of AtMPK1, AtMPK3 and AtMPK6 (Popescu et al., 2009).

138 To investigate the molecular mechanisms underlying the function(s) of AtTZF1, a Y-2-H screen to identify its protein-protein interactions was conducted by Zhang, L. Among the diverse classes of putative interacting partners revealed, three candidates with connections to stress response (AtMPK11, AtRDL2, AteIF2B) were further evaluated in this study. Based on the co-localization studies by Pomeranz et al., 2010a, several PB and SG markers were also evaluated for interactions with AtTZF1. While the full-length proteins of all these candidates failed to show a direct interaction in Y-2-H assays, the hypothesis that AtTZF1 may be regulated by the MAPK pathway was evaluated. An in silico analysis (conducted by Jang, J.-C.) revealed that AtTZF1 contains signature domains for MAP kinase- and phosphatase-based reversible phosphorylation. Based on the putative AtTZF1-AtMPK11 interaction revealed by the Y-2-H screen and the implications that AtMPK11 is involved in stress response including cold conditions (Menges et al., 2008), AtMPK11 appeared to be a good starting point to screen for the possible MAP kinase(s) that may link AtTZF1 to the stress-responsive MAPK signaling pathway in plants. However, an in vitro kinase assay failed to show phosphoryalation of AtTZF1 by AtMPK11 under the conditions employed. Alternative and/or improved assays are necessary to further evaluate these results.

3.3 Materials and Methods

3.3.1 Y-2-H screen

The Y-2-H screen that identified AtMPK11 (At1g01560), AtRDL2 (At4g16190) and AteIF2B (At3g07300) as putative interacting partners of AtTZF1 was conducted by Zhang, L., using the GAL4-activation domain (AD) Y-2-H cDNA library [Walker two- hybrid cDNA library, stock number CD4-10, Arabidopsis Biological Resource Center (ABRC)] as the prey. A full length AtTZF1 bait has been constructed in the BD-GAL4 Cam Phagemid vector (Stratagene, catalog number 235702) containing the GAL4-

139 binding domain (BD). Yeast strain PJ694A (James et al., 1996) has been used for the screen following standard protocols (Walhout and Vidal, 2001) and the bait was confirmed to lack self-activation activity prior to screening of interacting partners. The sequenced clones of AtMPK11, AtRDL2 and AteIF2B were partial, likely owing to the partial cDNA clones represented in the prey library.

3.3.2 Constructs

The Gateway-compatible entry clones were developed using standard cloning protocols by Invitrogen Inc. The cDNA clones for AtTZF1, AtRDL2 and AteIF2B available from ABRC (Stock numbers AtTZF1; U12422, AtRDL2; U12776 and AteIF2B; U67217) were used to amplify the coding regions via PCR. AtMPK11 was cloned by Zhang, L., from the pFL61 cDNA library (Minet et al., 1992). All inserts in the entry clones were confirmed by sequencing. Entry clones of AtDCP1 (At1g08370), AtDCP2 (At5g13570), AtAGO1 (At1g48410), AtXRN4 (At1g54490) and AtPABP8 (At1g49760) in pENTR3C vector (Invitrogen) were a kind gift of Pomeranz, M. Inserts in the entry vectors were moved to Gateway-compatible destination binary vectors; pGWB6 (N-terminal GFP fusion, CaMV35S promoter), pGWB9 (N-terminal His-tag fusion, CaMV35S promoter) and pGWB5 (C-terminal GFP fusion, CaMV35S promoter) by LR recombination reaction (Invitrogen). pGWB vectors were a kind gift of Tsuyoshi Nakagawa at the Shimane University, Japan. Free GFP construct in Gateway pK7FWG2 vector (Karimi et al., 2002) was a kind gift of Iris Meier laboratory.

For Y-2-H assays, entry clones described above were subjected to LR recombination reaction with the ProQuest pDEST22 or pDEST32 Gateway-compatible Y-2-H vectors (Invitrogen) according to the manufacturer’s protocol to create GAL4-Activation Domain (AD)- or GAL4-Binding Domain (BD)-fusion constructs respectively (Invitrogen handbook on ProQuest Two-Hybrid System with Gateway Technology, Series 10835, Version C, 2002).

140 To develop constructs containing AtTZF1 and AtMPK11 in the Tobacco mosaic virus RNA-based overexpression (pJL-TRBO) vector (Lindbo, 2007a; Lindbo, 2007b), AtTZF1 and AtMPK11 coding regions from pENTR/D-TOPO vector were subcloned with Not1 and AscI into a modified version of the pJL-TRBO vector in which the GFP tag has been removed and an AscI site has been added (a kind gift of Buckley, K. and Bisaro, D., Department of Molecular Genetics). Thereby, the configurations of the resulting proteins were HA2His6-AtTZF1 and HA2His6-AtMPK11. The clones were confirmed by sequencing.

3.3.3 Y-2-H assay

All tested plasmids pairs in Y-2-H assay were co-transformed into the yeast strain PJ694A (James et al., 1996) according to published protocols (Dohmen et al., 1991). Handling of yeast cultures and growth assays on media plates were conducted according to the ‘Clonetech Yeast Protocols Handbook, 1996’ (PT3024-1). Auxotrophic markers; leucine (–L for AD plasmids) and tryptophan (–T for BD plasmids) were used to select transformants. Growth of transformed yeast on media plates lacking histidine and adenine (–H,–A) or histidine alone (–H) indicated a positive protein-protein interaction.

3.3.4 Transient protein expression in Nicotiana benthamiana

The constructs in the pGWB binary destination vectors were transformed into the Agrobacterium tumefaciens strain GV3101. Transformants were selected on LB plates containing 50 µg/mL kanamycin, 10 µg/mL gentamicin, and 20 µg/mL rifampicin. Agrobacterium cultures containing each plasmid were co-inﬁltrated transiently into N. benthamiana leaves together with an Agrobacterium culture containing a construct that expresses the gene-silencing suppressor P19 as described in Zhao et al., 2006. Agrobacterium growth and inﬁltration conditions were adapted from Scofield et al., 1996 and Tang et al., 1996. In short, a preculture of Agrobacterium were inoculated into liquid

141 LB medium supplemented with 50 µg/mL kanamycin, 10 mM MES and 0.02 mM acetosyringone and grown overnight at 30°C. Cells were precipitated and resuspended in

10 mM MgCl2, 10 mM MES and 0.1 mM acetosyringone, to a final concentration of

OD600=1.0, incubated at room temperature ~3 h and injected into N. benthamiana leaves in a 1:1 mix with the Agrobacterium cell preparation expressing P19. Growth conditions of the infiltrated N. benthamiana plants varied in some experiments and are specified. Unless specified, post-infiltration growth conditions included; temperature 27°C, humidity 50%, regular watering and 16 h day/8 h night. Transient protein expression by infiltration with the A. tumefaciens strain C58C1 containing pJL-TRBO vectors were conducted as described by Lindbo, 2007a and Lindbo, 2007b.

3.3.5 Microscopy

Transiently transformed N. benthamiana epidermal cells were examined for GFP fluorescence and images were collected on a PCM 2000/Nikon Eclipse E600 confocal laser scanning microscope as described by Rose and Meier, 2001. For a gross comparative evaluation of GFP fluorescence as a measure of protein levels under varied conditions, infiltrated parts of the leaves showing the highest visual GFP fluorescence was observed with the low power lens (100X total magnification) of Nikon Eclipse E600 fluorescence microscope.

3.3.6 Antibodies

GFP-fusion proteins were detected with anti-GFP (Invitrogen; A11122), GST-fusion proteins with anti-GST (Sigma; G7781) and His-fusion proteins with anti-His (Santa Cruz Biotechnology; Sc-803) according to manufacturers’ protocols. AtTZF1 was detected with a polyclonal and affinity purified anti-peptide antibody developed in rabbit using the 5-21 amino acid fragment of AtTZF1 (Invitrogen, 1:1000 dilution).

142 3.3.7 Bioinformatics studies

Two motif search algorithms; Eukaryotic Linear Motif (Puntervoll et al., 2003) and Scansite 2.0 (Obenauer et al., 2003) were used to search for signature domains in AtTZF1 for MAPK signaling- based activation. (Analysis conducted by Jang, J.-C.)

3.3.8 Protein expression, purification and preparation for kinase assay

Kinase domain of SnRK1 (SnRK1-KD, ~42 kD) and SnRK1 kinase substrate (GST- SAMA, ~30 kD) were kind gifts of Mohannath, G. and Bisaro, D., Department of Molecular Genetics. Protein expression and purification protocols are described in

Mohannath, 2010. In short, these proteins were expressed as N-terminal HA2His6-fusion proteins using the Tobacco mosaic virus RNA-based overexpression (pJL-TRBO) vector in N. benthamiana transient expression system and purified by Ni-NTA column chromatography. During the preliminary trials, HA2His6-AtTZF1 and HA2His6- AtMPK11 proteins were also prepared by pJL-TRBO vector-based expression in N.

benthamiana following the same methodology. For HA2His6-AtTZF1 and HA2His6- AtMPK11 protein preparation using pJL-TRBO vector-based constructs in maize protoplasts, protoplast transformation and protein purification were carried out as described by Pomeranz et al., 2010a (protoplast associated work was carried out by Jang, J.-C.).

Myelin Basic Protein (MBP-Sigma M1891, ~18.4 kD) working solution was prepared at 1 µg/µl according to the manufacturer’s protocol. Four micrograms of MBP was added as the substrate to each kinase reaction mix. Bovine Serum Albumin (BSA-New England Biolabs B9001S, ~66.8 kD) was diluted from the 10 mg/ml commercial stock to use as standards for quantification of proteins.

143 For expression of GFP-AtMPK11 (~69.5 kD) and free GFP (~27 kD) proteins used for the kinase assays, Agrobacterium infiltration into N. benthamiana leaves for transient protein expression was conducted as described above. After preliminary trials using plants of varying maturity at infiltration, 3-week-old plants were selected for infiltration due to high GFP-fluorescence observed. Similarly, post-infiltration incubation was performed in cold room (temperature 5.5ºC, humidity 50%, regular watering, 16 h light/8 h dark) due to higher GFP florescence under these conditions compared to higher temperature conditions (temperature 27°C, humidity 50%, regular watering, 16 h light/8 h dark).

For purification of GFP-AtMPK11, anti-GFP magnetic beads (Miltenyi Biotech Inc.) were used according to the manufacturer’s protocol. Forty microliters of the bead slurry was used per 1 ml of plant powder. After the washes, bead-bound GFP-AtMPK11 was eluted under native conditions using 80 µl of 40 mM Tris-HCl pH 7.5 as the elution buffer. Elution was carried out by removing the column from the magnetic field after adding the native elution buffer. An equal amount of 80% glycerol was added prior to storage. This eluted extract containing GFP-AtMPK11 in the bead-bound form was used for the in vitro kinase assay. The glycerol added extract was denoted 1X concentration of GFP-AtMPK11. In order to increase the GFP-AtMPK11 concentration in the reaction mix in subsequent assays, the extract was stored without the addition of glycerol and denoted as 2X concentration of GFP-AtMPK11. For comparative assays, free GFP was purified at 2X concentration following the same methodology. 4X concentration of GFP- AtMPK11 and free GFP was achieved by adding twice the volume of 2X protein extract in the kinase reaction mix. For the purpose of evaluating the amount of magnetic bead- bound GFP-AtMPK11 and Free GFP on SDS-PAGE gels, protein elution under denaturing conditions was employed (Miltenyi Biotech Inc. protocols and adaptations thereof, as described in Figures 3.5 and 3.6).

144 Bacterially expressed and purified AtTZF1-GST fusion protein (~61.5 kD) (prepared according to standard protocols) was a kind gift of Qu, J. In short, the construct containing AtTZF1 in pDEST24 destination vector (Invitrogen) has been transformed into Escherichia coli BL21 (DE3) and protein expression was induced with isopropyl β- D-1-thiogalactopyranoside (IPTG). The AtTZF1-GST protein has been subsequently purified using Glutathion Sepharose 4B beads (GE Healthcare, 52-2303-00 AK) and eluted with reduced glutathione-containing buffer. To use AtTZF1-GST in the kinase assay, the reduced glutathione-containing buffer was subsequently exchanged with 40 mM Tris-HCl pH 7.5 via dialysis (Thermo Scientific, Slide-A-Lyser dialysis cassettes, 2160728, 3.5 kD cut off) in cold room according to the manufacturer’s protocol. The dialyzed AtTZF1-GST protein was subsequently concentrated using ultra centrifugal filter units (Millipore-Amicon, UFC 505008, 50 kD-cut off).

Transient expression of His-AtMPK11 in N. benthamiana leaves using pGWB9 vector was carried out in cold room and the tissues were harvested at 10 days post-infiltration (dpi). GFP-AtMPK11 was concurrently expressed as measure to evaluate protein expression by visualization of GFP fluorescence. Ni-NTA-based affinity purification was followed for His-AtMPK11 as described by Mohannath, 2010 for purification of

HA2His6-tagged proteins via pJL-TRBO-vector based expression, with elution using 250 mM imidazole-containing buffer.

3.3.9 Kinase assay

The in vitro kinase assay conditions standardized by Mohannath, 2010 were employed to assay phosphorylation of the previously established kinase-substrate pair SnRK-KD and GST-SAMA. The kinase reaction mix was prepared to contain the following final

concentrations: Tris-HCl pH 7.5 (50 mM), MgCl2 (10 mM), DTT (1 mM), Triton X-100 32 (1%), [γ- P] ATP (15 µCi, Perkin Elmer), HA2His6-SnRK1-KD (~15 ng) and HA2His6- GST-SAMA (~1 µg) in a total volume of 25 µl. The reaction mix was incubated for 40

145 min at 20ºC for the kinase reaction to occur, which was subsequently terminated by adding 3 µl of 0.5 M EDTA. The reaction mix was loaded to a SDS-10% polyacrylamide (w/v) gel of 1mm thickness with 12 µl of 3X protein loading buffer and subjected to eletrophoresis (i.e. SDS-Polyacrylamide Gel Electrophoresis : SDS-PAGE). Thus, the total volume of the reaction mix was maintained within the loadable volume per electrophoresis well of the gel apparatus (BioRad, mini-PROTEAN 165-8001), which is maximally ~44 µl/well for a 10 well, 1 mm-thick gel (BioRad mini-PROTEAN Tetra Cell 165-8001 Instruction manual), yet less than ~40 µl/well in most conditions to effectively load without overflow. Following Coomassie staining and destaining procedures, the gels were exposed to reusable storage phosphor screen (Kodak) for ~20 h. The radiographic images were acquired and analyzed using a Phosphorimager (Molecular Imager FX system-BioRad) and Quantity One 4.1.1 software (BioRad). Silver staining was performed after the radiographic exposure. Similarly, to evaluate MBP phosphorylation by SnRK-KD, the reaction mix was prepared containing 4 µg of MBP. For the phosphatase treatment, 10 units of Calf Intestinal Alkaline Phosphatase (CIP) (New England Biolabs, M0290S) were added to the kinase reaction mix prior to incubation.

Modifications were made to the kinase assay protocol explained above to improve the signal intensity when using GFP-AtMPK11 as the kinase. To facilitate detachment of GFP-AtMPK11 from the magnetic beads to enable movement of GFP-AtMPK11 through the SDS-PAGE gel (while the magnetic beads were retained in the well due to large size of beads), the post-incubation reaction mix was vortexed thoroughly for 2-3 min after adding the SDS-containing protein loading buffer. Other modifications such as the additional cations included as co-factors (at 10mM final concentration of each cation) and modifications in the volumes of components within the reaction mix are indicated in the results section and/or with figures. Kinase assays containing MBP (~18.4 kD) were conducted on SDS-12% polyacrylamide (w/v) gels.

146 3.4 Results

3.4.1 Evaluating putative AtTZF1 protein-protein interactions

3.4.1.1 Background

To identify molecular interactions of AtTZF1, a Y-2-H screen was conducted by Zhang, L. The functional categories of the putative interacting partners identified included regulators of transcription, translation and stress response. Three candidates with stress response-associate roles; AtMPK11, AtRDL2, AteIF2B were pursued in this study.

Among the twenty Arabidopsis MAPKs (dentoted AtMPK1-AtMPK20), AtMPK11 is an understudied plant MPK. However, functional implications could be derived from its closest relative, AtMPK4 which is well characterized (Kosetsu et al., 2010). AtMPK4 and AtMPK11 share 88.3% homology (Kosetsu et al., 2010), and are possibly activated by the same upstream MPK kinases (MPKKs) (Lee et al., 2008). They have recently been shown to be important for cytokinesis and AtMPK11 is transcriptionally elevated in atmpk4 mutant plants implicative of a compensatory mechanism (Kosetsu et al., 2010). These data together suggest that AtMPK11 and AtMPK4 may perform overlapping functions. AtMPK4 has been implicated in ABA signaling and is known to be involved in a range of abiotic and biotic stress tolerance-related signaling including drought, salt, cold, wounding and pathogen infestation (Colcombet and Hirt, 2008). Similarly, AtMPK11 has also shown to be transcriptionally upregulated in response to a variety of abiotic and biotic stresses including cold treatment (Menges et al., 2008). Therefore, it could be expected that AtMPK11 may also function in cold, drought and ABA signaling cascades as known for AtMPK4. Thus AtMPK11 could provide a functional link between AtTZF1 and the MAPK pathway that may underlie cold and drought tolerance of AtTZF1-OE plants (Lin et al., 2011) via an ABA-mediated pathway.

147 AtRDL2 is an uncharacterized RD (responsive to dehydration) cysteine-type proteinase- family protein (Bernoux et al., 2008). In cotton, the TZF protein GhZFP1 which enhances tolerance to biotic (fungus) and abiotic (salt) stresses was shown to interact with another RD-family protein; RD21A in a Y-2-H assay (Guo et al., 2009). However, the molecular mechanism of downstream action resulting from this interaction and leading to stress tolerance is unknown. Further, AtRD21A (At1g47128) is upregulated in response to water deficit stress (Yamaguchi-Shinozaki et al., 1992) similar to AtRDL2 (Bray, 2002). Therefore, it is intriguing to evaluate whether a bona fide interaction of functional significance in stress response exists between AtTZF1 and AtRDL2. eIF2B (eukaryotic initiation factor 2B) is a guanine nucleotide exchange factor involved in eukaryotic translation initiation. It converts eIF2-GDP into eIF2-GTP which is an initial step in the formation of the pre-initiation complex of translation. Translational regulation is an important regulatory point in gene expression including stress response. Eukaryotic eIF2B has been implicated in stress-mediated signaling (reviewed in Sonenberg and Hinnebusch, 2007). Phosphorylation of the α subunit of eIF2-GDP by stress-induced eIF2α kinase converts eIF2-GDP from a substrate to a competitive inhibitor of eIF2B, thereby negatively regulating translation in response to stress (Gomez et al., 2002; Sonenberg and Hinnebusch, 2007). This same phosphorylation event has been reported as a requisite initial signal for SG formation in response to environmental stresses (Anderson and Kedersha, 2008). Arabidopsis eIF2B-like proteins have been indicated, however have not been characterized and one of which is At3g07300 (denoted as AteIF2B in the current study). Evaluation of the putative interaction between AtTZF1 and AteIF2B may lead to novel insights on AtTZF1 function, possibly regarding stress- mediated translational repression.

148 3.4.1.2 Y-2-H assay to test interaction of AtTZF1 with AtMPK11, AtRDL2 and AteIF2B

The AtMPK11, AtRDL2 and AteIF2B clones interacting with the full-length AtTZF1 bait in the Y-2-H screen were of partial length. To evaluate these interactions as full-length proteins, Y-2-H assays were conducted using full length clones of AtMPK11, AtRDL2 and AteIF2B. AtTZF1 showed negative interaction with the full length clones (Figure 3.1).

3.4.2 Interactions of AtTZF1 with other putative candidates and self-interaction

3.4.2.1 Background

Several P-body (PB) and stress granule (SG) markers that showed co-localization with AtTZF1 as described in Pomeranz et al., 2010a were evaluated for interaction. PB markers thus tested were Decapping 2 (AtDCP2), Exoribonuclease 4 (AtXRN4) and Argonaute 1 (AtAGO1). The SG marker evaluated in the Y-2-H assay was Poly(A) Binding Protein 8 (AtPABP8). Decapping 1 (AtDCP1) which showed co-localization with AtTZF1 in PBs in further experiments (conducted by Jang, J.-C., unpublished data) was also tested.

AtDCP1 and AtDCP2 are components of the mRNA decapping complex important for mRNA degradation in the PBs (Fenger-Gron et al., 2005; Xu et al., 2006; Goeres et al., 2007). It has been proposed that interaction of TTP with components of the decapping machinery in the PBs may facilitate transfer of TTP-bound mRNAs from SGs to PBs for decay (Anderson and Kedersha, 2008). AtXRN4 is the functional orthologue of yeast and human XRN1, the exonuclease associated with the decapping complex that performs TZF- and ARE-based mRNA degradation in the 5’ to 3’ direction (Kastenmayer and Green, 2000; Souret et al., 2004; Potuschak et al., 2006; Bailey-Serres et al., 2009; Kim

149 et al., 2010). AtXRN4 has also been implicated in the degradation of several mRNAs via microRNA and small interfering RNA-mediated pathways (Gazzani et al., 2004; Souret et al., 2004). In human, both DCP2 and XRN1 have shown to interact with TTP in co- immunoprecipiation assays (Fenger-Gron et al., 2005; Lykke-Andersen and Wagner, 2005). AtAGO1, the widely studied AGO in plants, is a component (i.e. endonuclease) of RISC which is involved in microRNA-mediated mRNA degradation (Baumberger and Baulcombe, 2005; Axtell et al., 2007; Brodersen et al., 2008; Mi et al., 2008). Section 3.2.3 described the postulate that an interaction between TTP and AGO-eIF2C family members may direct microRNAs to the target mRNA. In human, the hTTP-hAGO2 interaction revealed was considered possibly responsible for this mechanism (Jing et al., 2005). In plant, AtAGO1 has shown to predominantly recruit microRNAs (Mi et al., 2008). AtPABP8 is postulated to play an equivalent role to mammalian PABP1 in SGs, which is conferring mRNA stability by protective binding to the Poly(A) tail, thereby deterring ribonuclease activity and regulation of translation (Afonina et al., 1998; Anderson and Kedersha, 2008; Pomeranz et al., 2010a).

Positive interactions of AtTZF1 with these components could reveal important details regarding molecular mechanisms underlying AtTZF1 functions. As PBs and SGs are induced under stress conditions, such interactions could also reveal stress response mechanisms involving AtTZF1. However, the N-terminal region of hTTP which is responsible for its interactions with the tested components of decapping and 5’-3’ exonuclease machinery including hDCP2 and hXRN1 (Lykke-Andersen and Wagner, 2005) is divergent in AtTZF1. Nevertheless, an alternative molecular mechanism may render interaction between AtTZF1 and these co-localized proteins possible. On the other hand, despite the low homology (18%) between hTTP and hBRF-1 in the N-terminal domain, they were able to perform mRNA decay activation in a similar manner (Lykke- Andersen and Wagner, 2005). Therefore it is possible that crucial structural/functional elements required for mRNA decay, including those important for protein-protein

150 interaction may be conserved in the N-terminus of hTTP, hBRF-1 and AtTZF1, though unrevealed by the analyses conducted in these studies.

A possible function for the putative CHCH Zn finger identified in the plant-unique region of AtTZF1 (see section 3.2.4) is unknown. Given the DNA-binding properties of TZF proteins, the common occurrence of dimerization in DNA-binding proteins and the involvement of C2H2 motifs in homo- and hetero-dimerization (Mello et al., 1996; Wang et al., 2008b; Pomeranz et al., 2010a; McCarty et al., 2003), it was attractive to test the possibility of homo-dimerization of AtTZF1, perhaps via the CHCH domain.

3.4.2.2 Y-2-H assay to test interactions of AtTZF1 with co-localized markers and self-interaction

Figure 3.2 shows that co-localized candidates failed to show an interaction with AtTZF1 under the assay conditions employed. Similarly, the test for a homotypic interaction of AtTZF1 (i.e. between AD-AtTZF1 and BD-AtTZF1) in full length clones also yielded a negative result.

3.4.3 Phosphorylation of AtTZF1 by AtMPK11

3.4.3.1 Background

In order to evaluate a possible link between the MAPK pathway and AtTZF1, AtMPK11 was tested as a MPK candidate for AtTZF1 in an in vitro phosphorylation assay. Although an interaction in full length clones could not be recapitulated for AtTZF1 and AtMPK11 in section 3.4.1.2, a phosphorylation relationship between a MAP kinase and its substrate does not necessarily require a strong and/or stable interaction (Sharrocks et al., 2000; Waas and Dalby, 2002; Menke et al., 2005). Also, it is possible that a bona fide in planta interaction may be hindered in the Y-2-H assay system when full length

151 AtMPK11 was tested. Evaluation of phosphorylation of AtTZF1 by AtMPK11 via an alternative approach compared to a protein-protein interaction assay was therefore pursued.

3.4.3.2 AtTZF1 contains signature domains for MAPK-based phosphorylation, and dephosphorylation

To gain more direct evidence regarding possible regulation of AtTZF1 by MAPK pathway, an in silico analysis of the AtTZF1 coding region was conducted by Jang, J.-C. It revealed that AtTZF1 contains signature domains for MAPK-mediated regulation via phosphorylation, and dephosphorylation (Figure 3.3). An N-terminal MAPK-docking site and several downstream putative serine phosphorylation sites were revealed, which is a spatial arrangement commonly found in substrates of MAPK-based regulation (Sharrocks et al., 2000; Andreasson et al., 2005). Further, a phosphatase-binding site is also revealed, suggesting reversible phosphorylation of AtTZF1 as a regulatory mechanism. Recapitulating hTTP’s mode of action in arsenite-induced stress response where phosphorylated hTTP interacts with 14-3-3, AtTZF1 also revealed a 14-3-3 interaction site (Figure 3.3).

3.4.3.3 Protein expression and purification for AtTZF1 and AtMPK11

Because MAP kinases are eukaryotic kinases that are themselves regulated by phosphorylation (Shirakabe et al., 1992; Whitmarsh and Davis, 1999), expression of AtMPK11 in a homologous system, particular in a plant-based system, was expected to be advantageous which could preserve its inherent kinase activity. Several plant-based protein production methods were tested to produce AtMPK11 and AtTZF1. These included pJL-TRBO vector-based transient expression and purification of HA2His6-

AtMPK11 and HA2His6-AtTZF1 from N. benthamiana (Lindbo, 2007a; Lindbo, 2007b) and maize protoplasts (protoplast-based expression was conducted by Jang, J.-C.), as well

152 as pGWB6 vector-based transient expression and purification of GFP-AtMPK11 and GFP-AtTZF1 from N. benthamiana. Post-infiltration incubation of N. benthamiana was carried out at 27°C, humidity 50%, regular watering and 16 h day/8 h night. AtTZF1 failed to show detectable expression by these methods when evaluated by GFP fluorescence and/or immunoblots from 1.5 to 12 dpi. However, when post-infiltration incubation of N. benthamiana expressing GFP-AtTZF1 was carried out in cold room at 5.5ºC, humidity 50%, regular watering, 16 h light/8 h dark, enhanced expression was observed 10 to 12 dpi (Figure 3.4). Similarly, the expression of GFP-MPK11 was enhanced when Agrobacterium-infiltrated N. benthamiana plants were incubated at 5.5ºC for 10 to 12 dpi. However, at 27°C AtMPK11 expression was observed at low levels when expressed by both pJL-TRBO and pGWB6 vectors. Immunoblot analysis showed that the protein expression level achieved for GFP-AtTZF1 in N. benthamiana transient expression at 5.5ºC was less than 20% compared to GFP-AtMPK11. In comparison, transgenic ~2-week-old Arabidopsis seedlings overexpressing AtTZF1-GFP showed higher AtTZF1 levels.

Accordingly, N. benthamiana plants transiently expressing GFP-AtMPK11 and subjected to post-infiltration incubation at 5.5ºC for 10 to 12 dpi were used for protein purification using anti-GFP magnetic beads. Both the input and the purified magnetic bead-bound GFP-AtMPK11 yielded a strong immunoblot signal at the expected position of ~69.5 kD when detected with the anti-GFP antibody (Figure 3.5A). However, the concentration of GFP-AtMPK11 obtained was low (~2 ng/µl), particularly considering the volume limitations in the kinase reaction mix (described in section 3.3.9). Low kinase concentrations have shown to be adequate to perform detectable substrate phosphorylation (an example shown in Figure 3.7A) likely dependant upon the kinase activity of the protein and optimized experimental conditions.

Purification of AtTZF1-GFP from transgenic Arabidopsis seedlings using magnetic anti- GFP beads yielded unacceptable low levels of purified AtTZF1-GFP to use as the

153 substrate in the kinase assay. Alternatively, His-fusion AtTZF1 was expressed in N. benthamiana using a pGWB9 vector-based construct following the protocol established including a 10 to 12 dpi cold-treatment at 5.5ºC. Nevertheless, Ni-NTA column-based purification yielded an unsatisfactory quality containing high background/co-purifying proteins. Therefore, bacterially expressed AtTZF1-GST (a kind gift of Qu, J.) and prepared further (see section 3.3.8 in Materials and Methods) was used as the substrate for the in vitro kinase assay. The GST-fusion AtTZF1 protein was detected by both anti- GST and anti-AtTZF1 antibodies (Figure 3.5C).

For comparative assays, GFP-AtMPK11 and free GFP were purified concurrently from 10 to 12 dpi cold-treated N. benthamiana according to the same procedure described above. Expected purification results were obtained as shown in Figure 3.6.

3.4.3.4 Optimization of the in vitro kinase- and phosphatase-assay protocols

To evaluate whether the in vitro kinase assay protocol employed by Mohannath, 2010 is successful in the current experimental context, a previously tested kinase-substrate pair (a kind gift of Mohannath, G. and Bisaro, D., Department of Molecular Genetics) was tested as a positive control. The protocol showed successful phosphorylation of the ~30 kD SnRK1 kinase substrate (GST-SAMA) by the kinase domain of SnRK1 (SnRK1-KD) and also showed SnRK1-KD (~42 kD) autophosphorylation as expected (Mohannath, 2010) (Figure 3.7A). Myelin Basic Protein (MBP) is known as a ‘universal kinase substrate’ due to its phosphorylation by a myriad of kinases (Romeis et al., 1999; Mayrose et al., 2004; Brock et al., 2010). To test the commercially available MBP as a kinase substrate to be used in subsequent experiments to optimize conditions for AtMPK11 activity, phosphorylation of MBP by SnRK1-KD was evaluated by in vitro kinase assay. MBP showed phophorylation by SnRK1-KD (Figure 3.7B). To optimize a dephosphorylation assay to further validate an in vitro phosphorylation result, incubation of Calf Intestinal Alkaline Phosphatase (CIP) with SnRK1-KD and MBP in the kinase assay was tested

154 (CIP treatment is described in Materials and Methods). Autoradiograph showed absence of the phosphorylation signals of MBP and SnRK1-KD with CIP treatment (Figure 3.7C).

3.4.3.5 Optimization of conditions for AtMPK11 kinase activity using MBP

To my knowledge, the protocol used for AtMPK11 purification using the anti-GFP magnetic beads of Miltenyi Biotech Inc. has not been reported previously to purify a kinase for a kinase activity-based assay. Therefore to assay whether the magnetic bead- bound GFP-AtMPK11 retains its kinase activity, an in vitro kinase reaction was conducted using MBP as the substrate. Figure 3.8A shows autoradiographic signals at the positions of MBP and GFP-AtMPK11 suggesting the ability of magnetic bead-bound GFP-AtMPK11 to phosphorylate MBP (A repeat experiment including a free GFP negative control is shown in Figure 3.10).

However, the level of MBP phosphorylation by GFP-AtMPK11 was extremely low compared to that of HA2His6-SnRK1-KD (Figure 3.8B) and required further optimization prior to conducting an assay to evaluate phosphorylation of AtTZF1-GST by GFP- AtMPK11. Higher concentration of AtMPK11 in the reaction mix (i.e. 4X AtMPK11 compared to 1X AtMPK11: see section 3.3.8) improved the MBP phosphorylation signal (Figure 3.9A). Incorporation of additional cations (i.e. Ca++ and Mn++) that could act as co-factors in enzymatic reactions (Maguire and Cowan, 2002) also improved the MBP phosphorylation signal in comparison to Mg++ alone (Figure 3.9B). However, it remains unclear if this improvement was brought on by an effect specific to Ca++ and/or Mn++ ions or instead by a general increase in the cation concentration.

To test if both the GFP-AtMPK11 protein extract and MBP suspension were required to yield the autoradiographic signal at the position of MBP, controls were tested with the elimination of each component. Figures 3.9A and C show that the observed MBP phophorylation signal required both the GFP-AtMPK11 extract and the MBP suspension

155 in the reaction mix, which supports that the signal likely pertained to phosphorylation of MBP by GFP-AtMPK11 extract and not a completely independent phosphorylation event in the reaction mix. At this point it still remained possible that another kinase in the GFP- AtMPK11 extract was able to phosphorylate MBP or a contaminant protein in the MBP preparation. However, the latter possibility seems highly unlikely due to the commercial purity of the MBP protein used. Figure 3.10 (lanes 4 and 5) which compares MBP phosphorylation signals between reaction mixes that contain GFP-AtMPK11 extract and free GFP extract shows that GFP-AtMPK11 and likely not a contaminating kinase in the GFP-AtMPK11 extract was responsible for the phosphorylation of MBP. This is key to using MBP to optimize conditions for GFP-AtMPK11 activity. To eliminate the remote possibility that a contaminant protein with affinity to AtMPK11 was responsible for MBP phosphorylation, a kinase-defective mutant version of GFP-AtMPK11 (for a possible methodology to construct a kinase-defective mutant, see Her et al., 1993) would serve as a suitable control.

3.4.3.6 Evaluation of phosphorylation of AtTZF1 by AtMPK11

To evaluate whether AtMPK11 is able to phosphoryate AtTZF1 in vitro, improved kinase assay conditions as described above were used. Magnetic bead-bound GFP-AtMPK11 and bacterially purified AtTZF1-GST were used for the assay. Autoradiograph in Figure 3.10 shows that GFP-AtMPK11 does not phosphorylate AtTZF1-GST at a detectable level (lane 1) although GFP-AtMPK11 is able to phosphorylate MBP when MBP is used at a lower concentration (~50%) compared to AtTZF1-GST (lane 4). Therefore, it appears that AtMPK11 may not be a bona fide kinase of AtTZF1 as determined by the in vitro assay system used here. If a kinase counterpart were known for AtTZF1, it could be used to evaluate the quality of the AtTZF1-GST used in the assay, which would serve as an important control. However, the protein band for AtTZF1-GST (~61.5 kD) in the Coomassie blot shows that the intact full-length protein was present in the reaction mix. Further, an unknown co-purifying protein of ~30kD in the AtTZF1-GST preparation

156 appears to be phophorylated by possibly a contaminating kinase in AtMPK11-GFP and Free-GFP protein extracts (Lanes 1, 2 and 3). This shows that the protein purification and preparation method used for AtTZF1-GST could yield kinase assay-compatible substrate proteins and also that the conditions utilized for the kinase assay in lane 1 was able to result phosphorylation. Thus, in addition to absence of interaction in full-length clones between AtMPK11 and AtTZF1 in the Y-2-H assay, a second means of preliminary data that fails to support a possible involvement of AtMPK11 in AtTZF1 activity was obtained.

3.5 Discussion and future directions

3.5.1 Interacting partners of AtTZF1

To elucidate the molecular mechanism(s) of AtTZF1 function, identifying its interacting partners is imperative. Direct interacting partners of AtTZF1, as well as components of functional complexes of AtTZF1 that may interact indirectly with AtTZF1 (i.e. via other proteins) are of significance for this purpose. This part of the study attempted to seek for direct interactions of AtTZF1. Y-2-H assays could primarily elucidate direct protein- protein interactions but may also include interactions that are mediated via yeast nuclear proteins. Interactions if identified via Y-2-H were intended to be tested via alternative methods to establish their ‘direct’ or ‘indirect’ nature.

Inability to recapitulate interactions of AtTZF1 with AtMPK11, AtRDL2 and AteIF2B in the Y-2-H assay using full length clones and alternative vector system could result due to several reasons. These may include conformational or stability issues of full length proteins and/or differences between the two vector systems. Particularly, the Gateway Y- 2-H system purposefully maintains low expression levels to improve the stringency of the result (Invitrogen handbook on ProQuest Two-Hybrid System with Gateway Technology,

157 Series 10835, Version C, 2002). Attempts to recapitulate these interactions in the same vector system used for the screen may be a suitable approach. Alternatively, the tested interactions including those with the co-localized markers could be tested via other interaction assay methods such as co-immunoprecipitation, BiFC and in vitro pull-down assays. Particularly an in vitro pull-down approach may help evaluate direct interactions of AtTZF1 thereby providing a higher level of information compared to the in vivo methods where additional proteins may mediate the interaction.

Further, an assay to evaluate AtTZF1 and At14-3-3 interaction particularly under various stress conditions including drought and cold stress in comparison to non-stress conditions could reveal useful information. It could help evaluate whether a situation similar to arsenite-induced and phosphorylation-based interaction of TTP with 14-3-3 seen in human (Stoecklin et al., 2004; Wilusz and Wilusz, 2004) would exist under the tested conditions in planta as well.

To identify components of in vivo functional complexes that contain AtTZF1, an immunoprecipitation/pull-down assay conducted using Arabidopsis plants would be particularly useful. Transgenic TAP tag-based immunoprecipitation may be a suitable option due to the enhanced specificity of the protocol to identify authentic candidates. In vivo immunoprecipitation screens under diverse environmental stress conditions including cold and drought could be particularly useful in interpreting the observed stress-tolerance phenomena of AtTZF1-OE plants. It may help identify functional complexes that are formed in response to particular stresses. Similarly, Y-2-H screens using cDNA libraries prepared from plants subjected to stress conditions of interest such as cold and drought [which may thereby represent the transcriptome induced under the particular stress conditions] could reveal specific interactions pertaining to each stress condition. In comparison to the immunoprecipitation approach in transgenic Arabidopsis plants, these Y-2-H screens may reveal a higher proportion of direct interacting partners.

158 Protein microarray-based screens (Popescu et al., 2007) using stress-induced proteomes would be expected to reveal direct interacting partners.

Particularly due to important implications of an interaction between AtTZF1 and AGO- eIF2C proteins in terms of the microRNA pathway, AGO-eIF2C family members could be screened for interaction. Because AtTZF1-AtAGO1 showed a negative interaction result in the Y-2-H assay conducted, co-immunoprecipitation experiments may be preferable in order to include candidates that interact via protein complex formation. Because in human, an interaction between hTTP and hAGO2 has been revealed, AtAGO2 seems worthwhile to be particularly tested for interaction with AtTZF1, despite that AtAGO2 is indicated in recruiting siRNAs rather than microRNAs in planta (Mi et al., 2008).

3.5.2 Protein expression with respect to temperature

Several widely used methods of protein expression was challenging particularly for AtTZF1. In our hands AtTZF1 has shown to be a highly labile protein with temperature sensitivity during protein extraction and immunoblot assays. During transient expression in N. benthamiana at 27°C (using both pJL-TRBO and pGWB6 vectors), possible instability of AtTZF1 may have caused failure to show detectable protein levels. Due to the implicated temperature sensitivity of AtTZF1, cold temperatures were tested to enhance its expression. Interestingly, enhancement of GFP fluorescence at 5.5°C was observed not only for GFP-ATTZF1 and GFP-AtMPK11 but also for several other fusion proteins expressed from pGWB5 and pGWB6 vectors (i.e. GFP-AtMPK3, GFP-AtMPK6, GFP-AtRDL2, AteIF2B-GFP and GFP-AtAGO1) as well as for free GFP expressed from pK7FWG2 (Appendix A and data not shown). To further test these results, a more precise quantification method to evaluate expression of GFP-fusion proteins by fluorescence detection [e.g. computational, spectrophotometric (Wydro et al., 2006; Sheludko et al., 2007)] together with an immunoblot-based analysis is required.

159 Certain trends in N.benthamiana transient protein expression at 5.5°C versus 27°C during 1.5- to 12-day post infiltration (dpi) period are noted. The tested proteins were GFP- AtTZF1, GFP-AtMPK11, GFP-AtMPK3, GFP-AtMPK6, GFP-AtRDL2, AteIF2B-GFP, GFP-AtAGO1 and free GFP. At 5.5°C (conditions described in 3.3.4), protein expression as determined by GFP fluorescence increased with the number of days post-infiltration for all tested proteins reaching the highest levels at 10 to 12 dpi. This contrasted with the expression trends under 27°C (conditions described in 3.3.8) where protein expression reached a peak ~2 to 5 dpi and diminished with passage of time. Thus, by 10 to 12 dpi, all tested proteins showed higher levels at 5.5°C compared to 27°C (Appendix A and data not shown). GFP-AtTZF1 and GFP-AtAGO1 which failed to show GFP fluorescence during the entire observation period under 27°C, showed moderate, but detectable protein expression 3 dpi at 5.5°C. At 3 dpi, all other tested proteins showed higher protein levels at 27°C compared to 5.5°C.

Because all these vectors contain the CaMV35S promoter for plant-based expression, it may be possible that in the cold protein expression from CaMV35S promoter occurs at a slower rate than at higher temperatures (e.g 27°C). However, in cold conditions, the expressed proteins may sustain with increased stability that leads to accumulation of the protein with time compared to 27°C. Temperature-effects on protein stability is known to depend upon the properties of a protein (Becktel and Schellman, 1987), which may explain why specifically GFP-AtTZF1 and GFP-AtAGO1 failed to accumulate detectable protein levels at 27°C. The ability to enhance protein levels may be useful for protein expression-related studies and protein-protein interaction assays. Particularly, assays involving AtTZF1 and AtAGO1 in the transient expression system in N. benthamiana may require elevated protein levels by cold-treatment due to undetectable protein levels at 27°C. Also, during efforts to identify conditions for N. benthamiana-based protein expression in literature for potential biolotechnological applications (e.g. plant-based production of therapeutic antibodies) (Sheludko et al., 2007), cold temperatures have not been tested.

160 3.5.3 AtTZF1, AtMPK11 and phosphorylation

The concentration of GFP-AtMPK11 obtained by the purification protocol was low, but was able to show detectable MBP phosphorylation. Although particular details are unavailable, Colcombet and Hirt, 2008 reports that for an unknown reason, detection of activity of MPKs other than MPK3, 4 and 6 has been challenging. Isolation of plant AtMPK11 that preserves its kinase activity, and identification of some conditions that facilitate MBP phosphorylation by AtMPK11 in this study could be helpful for evaluating AtMPK11 activity in future experiments including stress response studies (Doczi et al., 2007). The strikingly weaker MBP phosphorylation signal observed by GFP-AtMPK11

in comparison to HA2His6-SnRK1-KD could be due to several possible reasons. These include negative effects on the AtMPK11 kinase activity resulting from the purification methodology, magnetic bead-association of GFP-AtMPK11 or the GFP fusion per se, lower match between AtMPK11 and MBP as a kinase-substrate pair, lower inherent kinase activity of AtMPK11 and non-optimal assay conditions.

Absence of observable AtTZF1-GST phosphorylation provided a preliminary indication that GFP-AtMPK11 may not be a bona fide MAP kinase counterpart of AtTZF1. Further experiments are required to confirm this result. Particularly the ability of the purified AtTZF1-GST to be phosphorylated by an alternative MAP kinase would serve as a valuable positive control. Other possible MAP kinase candidates for AtTZF1 which include AtMPK1, AtMPK3 and AtMPK6 (Popescu et al., 2009) could be tested for this purpose. MBP could potentially serve as a substrate to optimize assay conditions for these MPKs as well (Popescu et al., 2009).

MBP showed detectable phosphorylation by GFP-AtMPK11 when present in the reaction mix at a lower concentration than AtTZF1-GST, which suggests that the loaded AtTZF1- GST amount may have been sufficient to show a detectable phosphorylation signal. However, if the match between GFP-AtMPK11 and AtTZF1-GST as a kinase-substrate

161 pair is lower compared to GFP-AtMPK11 and MBP, it may require higher activity of GFP-AtMPK11 and/or higher protein levels of AtTZF1-GST to impart a detectable level of phosphorylation. Particularly, improved expression and purification methodologies could be attempted to obtain higher concentration, purity and possibly improved kinase activity of AtMPK11. During production of active AtMPKs by transient expression in N. benthamiana for kinase assays by Popescu et. al., 2009, the upstream MPKK that phosphorylates the MPK of interest was also co-transformed. MPKK1, MPKK2 and MPKK6 which have shown to interact with AtMPK11 and AtMPK4 (Lee et al., 2008) could be co-transformed with AtMPK11 in protein production efforts to attempt improving kinase activity of AtMPK11. Also, the in vitro kinase assay protocol could be further optimized to obtain an enhanced phosphorylation signal. These include assay conditions that may enhance kinase activity of the MPK being tested and methodology that could accommodate higher component volumes particularly the kinase and substrate.

To test AtMPK-AtTZF1 phosphorylation reactions in vivo in a preliminary assay, the protoplast co-transformation methodology which has shown success with similar phosphorylation experiments (Doczi et al., 2007) could be employed. Currently, in the absence of a specific antibody that detects phosphoylated-AtTZF1, the differences in AtTZF1 phosphorylation status upon co-transformation with AtMPKs could be evaluated by an anti-phosphoprotein antibody (e.g. monoclonal anti-phosphoserine antibody. Sigma, catalog number P3430) or phosphoprotein gel stain (Pro-Q Diamond Phosphoprotein Gel Stain. Invitrogen, catalog number P33301) that detect phosphorylated proteins. If the baseline phosphorylation status of AtTZF1 assayed as a control in this experiment shows a positive result (i.e. baseline phosphorylation effectuated by endogenous kinases present in the protoplasts), it may support the results of the in silico analysis in section 3.4.3.2 that predicted a disposition of AtTZF1 to undergo phosphorylation. If successful, these phospho-protein detection methods could also be employed to evaluate in vivo phosphorylation status of AtTZF1 in AtTZF1-OE plants subjected various environmental conditions.

162

Figure 3.1: Y-2-H assay failed to show a direct interaction for AtTZF1- AtMPK11, AtTZF1- AtRDL2 and AtTZF1- AteIF2B using full-length clones. Gateway pDEST32 [GAL4-BD containing vector, auxotrophic marker: Leucine (L)] and pDEST22 [GAL4- AD containing vector, auxotrophic marker: Tryptophan (T)] were used to generate constructs. Construct pairs were co-transformed into PJ694A yeast strain. Growth results on Y-2-H media plates for several colonies containing each construct pair are shown. –L– T plates were used to select transformants. Growth on plates without hisidine or histidine and adenine (–H–A) indicates protein-protein interaction. Growth on –L–T plates indicates the presence of both constructs being tested. (A) AD-AtTZF1 shows negative results for interaction with BD-AtMPK11 and BD-AtRLD2. Similar results are obtained in the reciprocal direction (i.e. BD-AtTZF1 with AD-AtMPK11 and AD-AtRLD2). (B) AD-AtTZF1 shows negative results for interaction with BD-AteIF2B. Negative and positive controls show expected results. The positive control vector pair pEXP™22/RalGDS-w and pEXP™32/Krev1 was provided by the Invitrogen Y-2-H kit (Catalog #: PQ10001-01, 2005).

163 Figure 3.1

Colony number Colony number 1 2 3 4 1 2 3 4 BD-AtMPK11 AD-AtTZF1

BD-AtRDL2 AD-AtTZF1 BD AD-AtTZF1 BD-AtTZF1 AD-AtMPK11 BD AD-AtMPK11

AD-AtRDL2 BD-AtTZF1

BD AD-AtRDL2

BD-AtTZF1 AD

BD-AtMPK11 AD

BD-AtRDL2 AD BD AD

Y-2-H kit positive control

–L–T–H–A –L–T

Continued

164 Figure 3.1: Continued

Colony number Colony number 1 2 3 4 5 1 2 3 4 5 BD-AteIF2B AD-AtTZF1

BD AD-AtTZF1

Y-2-H kit positive control

BD-AteIF2B AD

BD AD –L–T–H –L–T

165

Figure 3.2: Y-2-H assay failed to show interactions between AtTZF1 and PB/SG markers, and AtTZF1 self-interaction. Gateway pDEST32 [GAL4-BD containing vector, auxotrophic marker: Leucine (L)] and pDEST22 [GAL4-AD containing vector, auxotrophic marker: Tryptophan (T)] were used to generate constructs. Construct pairs were co-transformed into PJ694A yeast strain. Growth results on Y-2-H media plates for 4 colonies containing each construct pair are shown. –L–T plates were used to select transformants. Growth on plates without histidine and adenine (–H–A) indicates protein-protein interaction. Growth on –L–T plates indicates the presence of both constructs being tested. BD-AtTZF1 showed negative results for interaction with the PB markers AD-AtDCP1, AD-AtDCP2, AD-AtXRN4 and AD-AtAGO1. Similarly BD-AtTZF1 failed to show an interaction with the SG marker AD-AtPABP8. No interaction was detected for BD-AtTZF1 + AD-AtTZF1 combination, thereby indicating an absence of a direct homo-dimerization. Negative and positive controls show expected results. The positive control vector pair pEXP™22/RalGDS-w and pEXP™32/Krev1 was provided by the Invitrogen Y-2-H kit (Catalog #: PQ10001-01, 2005).

166 Figure 3.2 Colony number Colony number 1 23 4 1 2 3 4 BD-AtTZF1 AD-AtDCP1 BD-AtTZF1 AD-AtDCP2 BD-AtTZF1 AD-AtXRN4 BD-AtTZF1 AD-AtAGO1 BD-AtTZF1 AD-AtPABP8 BD-AtTZF1 AD-AtTZF1 BD-AtTZF1 AD Y-2-H kit positive control BD AD-AtDCP1 BD AD-AtDCP2 BD AD-AtXRN4 BD AD-AtAGO1 BD AD-AtPABP8 BD AD-AtTZF1

BD AD –L–T–H–A –L–T

167 1 MMIGENKNRPHPTIHIPQWDQINDPTATISSPFSSVNLNSVNDYPHSPSPYLD 53 54 SFASLFRYLPSNELTNDSDSSSGDESSPLTDSFSSDEFRIYEFKIRRCARGRS 107 108 HDWTECPFAHPGEKARRRDPRKFHYSGTACPEFRKGSCRRGDSCEFSHGVFEC 160 161 WLHPSRYRTQPCKDGTSCRRRICFFAHTTEQLRVLPCSLDPDLGFFSGLATSP 213 214 TSILVSPSFSPPSESPPLSPSTGELIASMRKMQLNGGGCSWSSPMRSAVRLPF 266 267 SSSLRPIQAATWPRIREFEIEEAPAMEFVESGKELRAEMYARLSRENSLG 315

Figure 3.3: Sequence analysis revealed MAPK-docking, phosphorylation and phosphatase-binding sites in the AtTZF1 protein. Two motif search algorithms; Eukaryotic Linear Motif (Puntervoll et al., 2003) and Scansite 2.0 (Obenauer et al., 2003) identified a potential MAPK-docking site (in green), multiple phosphorylation sites (in red) and a protein phoshatase-interacting site (in blue) in AtTZF1. Underlined are the serine residues predicted to be phosphorylated. A potential 14-3-3 protein interacting site (RGRSHD) was also identified (in magenta).

Analysis and diagram construction were conducted by Jang, J.-C.

168 A B

Figure 3.4: Cold room conditions increased protein levels of N. benthamiana leaves expressing GFP-AtTZF1. Confocal micrographs of N. benthamiana leaves transiently expressing GFP-AtTZF1 at 11 days post-infiltration are shown under (A) 27°C and (B) 5.5°C (environmental conditions are described in 3.4.3.3.). GFP-AtTZF1 lacks an observable fluorescence signal under 27°C but is detected with high signal levels at 5.5°C. Scale bars: 100 µm.

169

Figure 3.5: Purification of GFP-AtMPK11 and AtTZF1-GST for in vitro kinase assay. (A) Silver staining and immunoblot results for GFP-AtMPK11 purification upon Agrobacterium-mediated transient expression in N. benthamiana leaves. Infiltrated plants were incubated in cold room for 11 days prior to tissue harvest. Purification was carried out using anti-GFP magnetic beads (Miltenyi Biotech Inc.). (B) Silver staining after purification shows GFP-AtMPK11 (~69.5 kD) with fewer co-purifying proteins compared to the numerous proteins visible in the input. Anti-GFP detects a strong band for GFP-AtMPK11 in both input and after purification. GFP-AtMPK11 was eluted under denaturing condition to prepare the ‘after purification’ SDS-PAGE gel and immunoblot. [i.e. a sample of GFP-AtMPK11 bound to anti-GFP magnetic beads collected via native elution was subsequently eluted under denaturing condition (Miltenyi Biotech Inc. protocol for in-column elution was modified for batch elution by microcentrifugation)]. (C) shows silver staining and immunoblot results of bacterially expressed and affinity purified AtTZF1-GST (a kind gift of Qu, J.) prior to buffer exchange and concentration. Expected band for AtTZF1-GST (~61.5 kD) was detected in both anti-AtTZF1 and anti- GST immunoblots overlapping with the AtTZF1-GST position in the silver-stained gel. The yield of purified product is low for both GFP-AtMPK11 and AtTZF1-GST proteins.

170 Figure 3.5

A B

kD 75 kD 75

50 50

Silver stain α-GFP Silver stain α-GFP

Input After purification

C kD kD kD 75 75 75

50 50 50

37 37 37 Silver stain α-GST α-AtTZF1

171

GFP-AtMPK11 Free GFP GFP-AtMPK11 Input Input Free GFP Purified Purified Input Input Purified Purified

kD kD 75 75

50 50 37 37

25 25 α-GFP Silver stain

Figure 3.6: GFP-AtMPK11 and free GFP were prepared concurrently for comparative in vitro kinase assays. Silver staining and immunoblot results of GFP- AtMPK11 and free GFP purified from Agrobacterium-infiltrated and 11 day cold room- treated N. benthamiana, using anti-GFP magnetic beads (Milteny Biotech Inc.). Magnetic bead-bound proteins were eluted from columns under native conditions to be used in the in vitro kinase assays (described in Materials and Methods). For the preparation of SDS- PAGE gel and immunoblot shown here, a sample of proteins bound to anti-GFP magnetic beads collected via native elution was subsequently eluted under denaturing condition (Miltenyi Biotech Inc. protocol for in-column elution was modified for batch elution by microcentrifugation). The positions for GFP-AtMPK11 (~69.5 kD) and free GFP (~27 kD) proteins are denoted by red and blue arrows, respectively. Purification yielded GFP- AtMPK11 and free GFP proteins at low concentration. Contaminating proteins are seen in the silver stained gel in both ‘GFP-AtMPK11-purified’ and ‘Free-GFP-purified’ lanes particularly ~55 kD and ~30 kD.

172

Figure 3.7: in vitro kinase assay detected phosphorylation/de-phosphorylation events under the experimented conditions. A previously established kinase (SnRK1-KD, ~42 kD ) and substrate (SnRK1-KD substrate GST-SAMA, ~30 kD ) pair was tested in an in vitro kinase assay. (A) The autoradiograph shows phosphorylation of the SnRK1-KD substrate by SnRK1-KD and SnRK1-KD autophosphorylation under the conditions employed. (B) The universal kinase substrate Myeline Basic Protein (MBP, ~18.4 kD ) shows phosphorylation by SnRK1-KD. Also SnRK1-KD autophosphorylation is seen. (C) Calf Intestinal Alkaline Phosphatase (CIP) abolishes the phosphorylation signal of MBP and SnRK1-KD (right lane). Silver and Coomassie stained gels show loading of the proteins. Exposure time: ~20 h.

173 Figure 3.7

A C SnRK1-KD + SnRK1-KD MBP + + MBP CIP 50kD SnRK1-KD

SnRK1-KD

Autoradiograph

SnRK1 substrate Silver stain MBP Autoradiograph 15kD MBP Coomassie

SnRK1-KD

MBP

Coomassie Autoradiograph

174

Figure 3.8: Magnetic bead-bound GFP-AtMPK11 is able to phosphorylate MBP. The elute containing GFP-AtMPK11 purified using anti-GFP magnetic beads (Milteny Biotech Inc.) was tested for the ability to phosphorylate MBP in an in vitro kinase assay. (A) GFP-AtMPK11 can phosphorylate MBP. [A repeat experiment including a free GFP negative control is shown in Figure 3.10] (B) A comparison between the signal strength of MBP phosphorylation by GFP-AtMPK11 and HA2His6-SnRK1-KD. Similar amounts of the two kinases (~15 ng) and 1X concentration of GFP-AtMPK11 [described in Materials and Methods] were used. The autoradiographic signal for MBP- phosphorylation by HA2His6-SnRK1-KD is extremely strong compared to the MBP- phosphorylation signal by GFP-AtMPK11. Coomassie-stained gels show protein loading. Exposure time: ~20 h.

175 Figure 3.8

MBP

GFP- HA2His6- AtMPK11 + SnRK1-KD + MBP MBP

MBP Autoradiograph Lower gain settings

Autoradiograph MBP Higher gain settings

MBP Coomassie

176

Figure 3.9: MBP phosphorylation by AtMPK11 was improved via modifications of the in vitro kinase assay protocol. (A) MBP phosphorylation is improved by higher concentration of GFP-AtMPK11 in the reaction mix. 1X and 4X GFP-AtMPK11 concentrations are described in Materials and Methods (section 3.3.8). (B) Addition of cations Ca++ and Mn++ that may act as co-factors improved the MBP phosphorylation signal by GFP-AtMPK11. (C) Further evaluation whether an independent phosphorylation reaction in the reaction mix (i.e. unrelated to GFP-AtMPK11 or MBP) could be responsible for the radiographic signal at the position of MBP. MBP- phosphorylation is seen in GFP-AtMPK11 + MBP, which is above the background signal level visible in the lane containing GFP-AtMPK11 alone. Coomassie-stained gels show protein loading. Exposure time: 2 days.

177 Figure 3.9

A 1X 4X MBP only GFP-AtMPK11 GFP-AtMPK11 MBP Autoradiograph

MBP Coomassie

B MgCl2+ MgCl2 MnCl2+ only CaCl2

Autoradiograph MBP

MBP Coomassie

C GFP-AtMPK11 + MBP GFP- AtMPK11

Autoradiograph MBP

MBP Coomassie

178 GFP- AtMPK11 GFP- + AtMPK11 Free GFP AtTZF1- GFP- + + GST AtMPK11 Free GFP AtTZF1- AtTZF1- + + + GST GST CIP MBP MBP 1 2 3 4 5

kD 75 AtTZF1-GST 50 37 Autoradiograph 25 20 MBP

75 AtTZF1-GST 50 37 Coomassie 25 20 MBP

Figure 3.10: AtMPK11 phosphorylates MBP, but not AtTZF1, in the in vitro kinase assay. The variables in the reaction mix for each lane are as indicated. Top panel shows the radiographic signals. Bottom panel shows protein loading with Coomassie staining. AtTZF1-GST (~61.5 kD) fails to show a radiographic signal in lane 1, but MBP (~18.4 kD) shows a radiographic signal in lane 4. Both reactions contained GFP-AtMPK11 as the kinase and MBP is much lower in concentration than AtTZF1-GST, as seen in the Coomassie stained gel. MBP phosphorylation is negligible when free GFP (purified following the same protocol as GFP-AtMPK11) was used as the kinase in lane 5. A likely contaminant protein of ~30 kD is seen phosphorylated in lanes 1 and 2, which does not show the radiographic signal in lane 3 when CIP was contained in the reaction mix. Lanes 1, 2 and 3 in the Coomassie stained gel show thick lower bands (~22-26 kD) corresponding to degradation products of AtTZF1-GST. Exposure time: 4 days.

179 Appendix A: GFP-AtAGO1 and GFP-AtMPK3 expression at 27°C and 5.5°C

27ºC 27ºC 5.5ºC 3 days post-infiltration 11 days post-infiltration 11 days post-infiltration A B C

GFP- AtAGO1

D E F

GFP- AtMPK3

Expression of GFP-AtAGO1 and GFP-AtMPK3 at 27°C and 5.5°C. Agrobacterium- infiltrated N. benthamiana leaves were observed for GFP fluorescence. GFP-AtAGO1 expression is undetectable at (A) 3 dpi and (B) 11 dpi at 27°C [At 3 dpi, very faint GFP- AtAGO1 expression was observable at 5.5°C (data not shown)]. (C) At 11 dpi at 5.5°C GFP-AtAGO1 shows detectable expression. In comparison, reasonably high expression of GFP-AtMPK3 was detected at (D) 3 dpi at 27°C. GFP-AtMPK3 expression diminished to barely detectable levels at (E) 11 dpi at 27°C. (F) At 11 dpi at 5.5°C, GFP- AtMPK3 shows very high expression. All images were collected at the same microscope settings. Scale bars: 100 µm.

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