Identification of Plant SUN Domain-Interacting Tail Proteins and Analysis of Their Function in Nuclear Positioning

DISSERTATION

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

Xiao Zhou, M. S.

Graduate Program in Plant Cellular and Molecular Biology

The Ohio State University

2013

Dissertation Committee:

Professor Iris Meier, Advisor

Professor Biao Ding

Professor Stephen Osmani

Professor R. Keith Slotkin

Xiao Zhou

2013

Abstract

The nuclear envelope (NE) is a double membrane system consisting of an inner nuclear membrane (INM) and an outer nuclear membrane (ONM). Studies in opisthokonts revealed that the two membranes are bridged by protein complexes formed by the INM

Sad1/UNC-84 (SUN) proteins and the ONM Klarsicht/ANC-1/Syne-1 homology

(KASH) proteins. These SUN-KASH NE bridges are usually linkers of the nucleoskeleton to the cytoskeleton (LINC) conserved across eukaryotes. LINC complexes are key players in multiple cellular processes, such as nuclear and chromosomal positioning and nuclear shape determination, which in turn influence the gametogenesis and several aspects of development. Although these cellular processes have long been also known in plants, no KASH proteins are encoded in the plant genomes.

I identified WPP domain interacting proteins (WIPs) as the first plant KASH protein analogs. WIPs are plant-specific ONM proteins that redundantly anchor Ran GTPase activating protein (RanGAP) to the NE. Arabidopsis thaliana WIPs (AtWIPs) interact with Arabidopsis thaliana SUN proteins (AtSUNs), which is required for both AtWIP1 and AtRanGAP1 NE localization. In addition, AtWIPs and AtSUNs are necessary for maintaining the elongated nuclear shape of Arabidopsis epidermal cells. These data

ii provide a novel function of the SUN-dependent NE bridges, suggesting that a functionally diverged SUN-dependent NE bridge is conserved beyond the opisthokonts.

Further analysis of AtWIPs and its binding partner Arabidopsis thaliana WPP domain- interacting tail anchored proteins (AtWITs) revealed that they are localized to the vegetative NE in pollen and play a role in the nuclear movement of the vegetative nucleus (VN) during pollen tube growth. Loss of AtWIPs or AtWITs resulted in impaired

VN movement and inefficient sperm-cell-to-ovule delivery. AtWIPs and AtWITs are the first genes assigned to nuclear movement during pollen tube growth.

WIPs have no similarity to known opisthokont KASH proteins, except for a C-terminal transmembrane domain, followed by a short SUN-domain interacting tail (SIT) domain terminating in a conserved four-amino-acid motif critical for the SUN-WIP interaction.

This conserved pattern was used to computationally search for candidate SIT proteins. As a second criterion, I asked for conservation of the rule in most homologs of a protein family. I identified 10 new potential SIT protein families, four of which were verified for their SUN-interaction-dependent NE localization. One SIT protein, Arabidopsis thaliana

AN NEMO 1, was shown to be associated with F-actin and involved in guard cell nuclear central anchorage. This study dramatically expands the number of SIT proteins and implies an independent evolution of SUN-dependent NE-bridges after the opisthokont- plant separation.

iii

Dedication

This document is dedicated to the nerds who still have a dream in science.

Acknowledgments

I was extremely fortunate to get the opportunity to work under the instruction of my advisor, Professor Iris Meier who offered me a free-thinking research environment and guided me to an interesting research field. Without her continuous guidance, support and motivation, this dissertation would not have been possible.

I am grateful to all my committee members, Professor Biao Ding who always challenged my critical thinking, Professor Steve Osmani who was very strict on my logic and data interpretation, and Professor Keith Slotkin who provided helpful advice and support on the project described in CHAPTER 3.

I would also like to thank all my lab-mates through the years: Dr. Jelena Brkljacic, Dr.

Sowmya Venkatakrishnan, Dr. Sivaramakrishnan Muthuswamy, Dr. Thushani Rodrigo-

Dr. Peiris, Dr. Joanna Boruc, Dr. Mintu Desai, Dr. Chris DeFraia, Alex Tough, Anna

Newman, Norman Groves, and other undergraduate assistants. I am grateful to their warmth, encouragement, and helpful discussion. I also want to thank Emily Yoders-Horn and Joan Leonard who took good care of my plants and set up the delightful greenhouse pond.

I am sincerely thankful to Dr. Katja Graumann and Professor David Evans for the delightful collaboration on the SUN-WIP project (CHAPTER 2). I want to thank Dr.

Xianfeng Xu who left detailed information on his materials which saved me considerable time on this project.

I want to thank my parents who always pointed out the negative side of my life as a researcher. This let me understand how to communicate with people outside of my research field.

In the end, I want to thank Dr. Yiyi Zhang, without her encouragement, I would not have come to study in the Ohio State University.

Vita

2004 B.S. Bioengineering, East China University of Science and Technology

2007 M.S. Biochemistry and Molecular Biology, Fudan University

2007 to present Graduate Researching Associate, Department of Molecular Genetics, The Ohio State University

Publications

Zhou, X., J. Boruc, and I. Meier. 2013. The Plant Nuclear Pore Complex — The Nucleocytoplasmic Barrier and Beyond. In Annual Plant Reviews. John Wiley & Sons Ltd. 57-91. Zhou, X., and I. Meier. 2013. How plants LINC the SUN to KASH. Nucleus. 4:206-215. Zhou, X., K. Graumann, D.E. Evans, and I. Meier. 2012. Novel plant SUN-KASH bridges are involved in RanGAP anchoring and nuclear shape determination. Journal of Cell Biology. 196:203-211. Boruc, J., X. Zhou, and I. Meier. 2012. Dynamics of the Plant Nuclear Envelope and Nuclear Pore. Plant Physiology. 158:78-86. Meier, I., X. Zhou, J. Brkljacic, A. Rose, Q. Zhao, and X.M. Xu. 2010. Targeting proteins to the plant nuclear envelope. Biochemical Society Transactions. 38:733-740.

Fields of Study

Major Field: Plant Cellular and Molecular Biology

vii

Table of Contents

Abstract ...... ii

Dedication ...... iv

Acknowledgments...... v

Vita ...... vii

Publications ...... vii

Fields of Study ...... vii

List of Tables ...... xv

List of Figures ...... xvi

CHAPTER 1 Essential function of SUN-KASH complexes in opisthokont nuclear positioning and the importance of plant nuclear positioning ...... 1

1.1 Abstract ...... 2

1.2 Introduction to SUN and KASH proteins ...... 3

1.3 Function of opisthokont SUN and KASH proteins ...... 4

viii

1.3.1 Nuclear positioning mediated by opisthokont LINC complexes...... 4

1.3.2 Chromosome movement mediated by opisthokont LINC complexes ...... 7

1.3.3 Nuclear pore complex assembly regulated by SUN1 ...... 7

1.4 Plant SUN proteins and their functions ...... 8

1.4.1 Plant SUN proteins ...... 8

1.4.2 Possible function of plant SUN proteins in mitosis and meiosis ...... 9

1.5 Nuclear positioning in plants...... 10

1.5.1 Nuclear movement of pollen tubes ...... 11

1.5.2 Nuclear movement of root and leaf hair ...... 13

1.5.3 Nuclear positioning during asymmetric cell divisions ...... 14

1.5.4 Root nuclear movement responding to microbes ...... 15

1.5.5 Nuclear positioning responding to physical stimuli ...... 16

1.6 Plant KASH analogs involved nuclear positioning ...... 17

CHAPTER 2 Novel plant nuclear envelope bridges are involved in RanGAP anchoring and nuclear shape determination ...... 28

2.1 Abstract ...... 29

2.2 Introduction ...... 30

2.3 Material and Methods...... 32

2.3.1 Plant materials ...... 32

2.3.2 Constructs ...... 32

2.3.3 Generation of transgenic plants ...... 33

2.3.4 Co-immunoprecipitation experiments ...... 33

2.3.5 DAPI staining and nuclear length measurement ...... 34

2.3.6 Root hair and trichome nuclear positioning Assay ...... 35

2.3.7 Confocal microscopy and FRAP ...... 35

2.4 Results ...... 35

2.4.1 Identification of AtWIPs as ONM AtSUN-interacting partners ...... 35

2.4.2 AtSUNs interact with AtWIP1 through their SUN domain on the NE ...... 37

2.4.3 AtSUN1 affects the mobility of AtWIP1 at the plant NE (data generated by

Katja Graumann) ...... 38

2.4.4 AtSUN1 and AtSUN2 are required to anchor AtWIP1 to the NE ...... 40

2.4.5 AtSUNs are required for AtRanGAP1 NE localization ...... 41

2.4.6 AtSUNs and AtWIPs are required for maintaining an elongated nuclear shape

in epidermal cells ...... 43

2.5 Discussion ...... 46

2.5.1 Interactions between AtSUN and AtWIP ...... 46

2.5.2 Function of the AtSUN-AtWIP complex in RanGAP NE anchoring ...... 46

2.5.3 Function of the AtSUN-AtWIP complex in nuclear shape determination ...... 48

CHAPTER 3 Efficient plant sperm cell delivery requires vegetative nuclear movement mediated by WPP domain-interacting tail-anchored proteins ...... 61

3.1 Abstract ...... 62

3.2 Introduction ...... 63

3.3 Material and Methods...... 67

3.3.1 Plant materials ...... 67

3.3.2 Arabidopsis cDNA synthesis ...... 67

3.3.3 Constructs ...... 68

3.3.4 Generation of transgenic plants ...... 70

3.3.5 Co-immunoprecipitation experiments ...... 70

3.3.6 Hoechst 33342 staining and nuclear shape measurement ...... 71

3.3.7 In vitro pollen germination and Alexander staining ...... 72

3.3.8 Pollen tube confocal microscopy, length measurement, and kymograph

generation ...... 72

3.4 Results ...... 73

3.4.1 WIT and WIP family proteins are required for full seed set ...... 73

3.4.2 Loss of WITs and WIPs leads to defects of pollen fertility ...... 73

3.4.3 WIT1 and WIP1 are located at the vegetative NE in pollen grains ...... 74

3.4.4 WIT and WIP family proteins are required for proper nuclear movement in

pollen tubes ...... 75

3.4.5 Loss of WIT proteins impaired the SC-to-ovule delivery ...... 78

3.4.6 VN movement is not mediated through the proposed SUN-WIP-WIT-myosin

XI-I complex ...... 79

3.5 Discussion ...... 81

3.5.1 The importance of MGU ...... 81

3.5.2 Driving forces of the MGU ...... 81

3.5.3 Difference between nuclear movement in vegetative tissues and pollen tubes 83

CHAPTER 4 DORY: an effective algorithm to discover a multitude of novel plant SUN domain-interacting tail proteins ...... 94

4.1 Abstract ...... 95

4.2 Introduction ...... 96

4.3 Material and Methods...... 99

4.3.1 DORY and BLASTP search ...... 99

4.3.2 Protein model correction based on the “transmembrane domain (TMD)-pSIT”

architecture ...... 100 xii

4.3.3 Plant materials ...... 100

4.3.4 Constructs ...... 101

4.3.5 Agrobacterium transformation, N. benthamiana transient expression, and

Arabidopsis stable transformation ...... 102

4.3.6 Co-immunoprecipitation experiments ...... 103

4.3.7 Confocal microscopy ...... 103

4.3.8 Nuclear position measurement in guard cells ...... 104

4.4 Results ...... 105

4.4.1 Identify Arabidopsis thaliana SUN-interacting protein candidates ...... 105

4.4.2 Verify the subcellular localization of SUN-binding protein candidates ...... 107

4.4.3 AtNEMO1, AtNEMO2, AtNEMOL, AtNEMOS, and MsNEME1 interact with

AtSUN1 and AtSUN2 ...... 108

4.4.4 The NE Localization of AtNEMO1, AtNEMO2, AtNEMOL, AtNEMOS, and

MsNEME1 depends on AtSUN ...... 111

4.4.5 The armadillo-repeat (ARM) domain of AtNEMO1 is associated with F-actin

...... 112

4.4.6 AtNEMO1 has a specific expression and localization pattern ...... 113

4.4.7 AtNEMO1 is required for proper nuclear anchorage in guard cells ...... 114

4.5 Discussion ...... 116 xiii

4.5.1 DORY as a useful tool to identify potential SIT proteins ...... 116

4.5.2 Diversity of plant SIT proteins and opisthokont KASH proteins ...... 117

4.5.3 Function of AtNEMO1 in guard cell nuclear anchorage ...... 118

4.5.4 The LINC function of plant SUN-KASH NE bridges ...... 120

CHAPTER 5 Perspectives ...... 140

5.1 Regulation of nuclear shape and its function ...... 141

5.2 Potential motor proteins in pollen nuclear movement ...... 142

5.3 Players in nuclear positioning other than SIT proteins ...... 142

5.4 Function of AtNEMO1 and AtNEMO2 ...... 143

5.5 Potential additional, unidentified SIT proteins ...... 144

References ...... 145

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List of Tables

Table 1.1 Known SUN-KASH/SIT pairs from different organisms, their cytoplasmic partners, and their known or proposed functions...... 24

Table 4.1 Primers used for cloning...... 137

Table 4.2 Primers used for genotyping nemo1-1, nemo1-2, nemo1-3, nemo2-1, and nemo2-2...... 139

List of Figures

Figure 1.1 SUN-KASH/SIT bridges across the NE in different organisms...... 20

Figure 1.2 Computed three-dimensional model of the SUN domain of AtSUN1...... 21

Figure 1.3 Amino acid sequence alignment of plant SUN domains with the SUN domain of HsSUN2...... 23

Figure 2.1 Structural and sequence similarity between KASH domains and the PNS tail of AtWIP1...... 50

Figure 2.2 Characterization of AtSUN-AtWIP interactions...... 51

Figure 2.3 FRAP analysis of the interaction between AtWIP1 and AtSUN1...... 52

Figure 2.4 AtSUNs are required for targeting AtWIP1 and AtRanGAP1 to the NE...... 53

Figure 2.5 Nuclear shape change in epidermal cells of sun1-KO sun2-KD and wip1-1 wip2-1 wip3-1 plants...... 54

Figure 2.6 Plant-specific conserved residues in plant SUN domains revealed by the alignment of SUN domains from different species...... 56

Figure 2.7 AtWIP1 and AtSUN localization and characterization of sun1-KO sun2-KD. 58

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Figure 2.8 Nuclear position in root hairs and trichomes is not affected in sun1-KO sun2-

KD, and wip1-1 wip2-1 wip3-1...... 60

Figure 3.1 Reduction in seed production based on loss of WITs and WIPs...... 84

Figure 3.2 WIT1 and WIP1 subcellular localization in pollen grains and pollen tubes. .. 85

Figure 3.3 Wit1-1 wit2-1, wip1-1 wip2-1 wip3-1, and wifi pollen conditions...... 86

Figure 3.4 Nuclear movement in pollen tubes is affected in wit1-1 wit2-1, wip1-1 wip1-1 wip2-1 wip3-1, and wifi...... 88

Figure 3.5 Distance between the leading nucleus and the tip of the growing pollen tube after the nucleus entered the pollen tube...... 89

Figure 3.6 Nuclear position after 8 h of semi-in-vitro pollen tube germination...... 90

Figure 3.7 Wit1-1 wit2-1 has SC delivery defects...... 91

Figure 3.8 Root hair and trichome nuclear shape is regulated by WIT2...... 92

Figure 3.9 WIT2* interact with and WIP1, WIP2, and WIP3...... 93

Figure 4.1 Development of DORY Algorithm ...... 123

Figure 4.2 Amino acid sequence alignment of the C-terminal domains of predicted plant

SIT proteins...... 125

Figure 4.3 Amino acid sequence alignment of the C-terminal domains of predicted SIT proteins not verified in this study...... 127

Figure 4.4 Subcellular localization of predicted plant SIT proteins...... 128

xvii

Figure 4.5 Co-IP analysis of AtNEMO-AtSUN interactions...... 130

Figure 4.6 Co-IP analysis of the AtNEMOL/AtNEMOS/MsNEME1-AtSUN interactions.

...... 131

Figure 4.7 F-actin association of AtNEMO11-308 (containing the ARM domain) ...... 132

Figure 4.8 Expression pattern of AtNEMO1 and AtNEMO2...... 134

Figure 4.9 AtNEMO1 is involved in guard cell central nuclear anchorage...... 135

Figure 4.10 Phenotype comparison between nemo1 and nemo2 mutants and wild type.136

xviii

CHAPTER 1

Essential function of SUN-KASH complexes in opisthokont nuclear positioning and

the importance of plant nuclear positioning

1.1 Abstract

Sad1/UNC-84 (SUN) proteins and the ONM Klarsicht/ANC-1/Syne-1 homology

(KASH) proteins. These SUN-KASH complexes are usually linkers of the nucleoskeleton to the cytoskeleton (LINC) and are conserved across eukaryotes. LINC complexes are key players in multiple cellular processes, such as nuclear and chromosomal positioning and nuclear shape determination, which in turn influence the gametogenesis and several aspects of development. Although these cellular processes have long been known in plants, no KASH proteins are encoded in the plant genomes. In this chapter, the composition of opisthokont LINC complex and its function in nuclear position will be reviewed. In comparison, the importance of plant nuclear positioning will be addressed, and the dissertation research on identification and functional analysis of plant KASH analogs will be introduced.

1.2 Introduction to SUN and KASH proteins

The founding member of SUN proteins is Caenorhabditis elegans UNC-84. The C- terminal domain of UNC-84 shares homology with two human proteins and

Schizosaccharomyces pombe Sad1, a component of the spindle pole body (Malone et al.,

1999). This domain was subsequently found to interact with a C-terminal domain conserved in a series of ONM proteins including Klarsicht, ANC-1, and Syne (also known as Nesprin) (Starr and Han, 2002). Therefore, the two interacting domains were named the UNC-84/Sad1 (SUN) domain (Malone et al., 1999) and Klarsicht/ANC-

1/Syne-1 homology (KASH) domain (Starr and Han, 2002), respectively. SUN proteins are typically located at the INM with their C-terminal SUN domain positioned in the perinuclear space (PNS) (Figure 1.1 and see below). KASH proteins are type-II transmembrane proteins or tail-anchored proteins at the ONM harboring a PNS KASH domain. The KASH domain terminates in a short tail (~30 amino acids) typically ending with a PPPX (X represents any amino acid) motif that is essential for interacting with the

SUN domain. The SUN-KASH complexes bridge their binding partners across the NE

(Figure 1.1). In the nucleoplasm, the N-termini of many SUN proteins interact directly or indirectly with nuclear lamins, intermediate filament proteins located underneath the

INM, which are considered a component of the nucleoskeleton (Stuurman et al., 1998).

At the cytoplasmic side, KASH proteins are linked to motor proteins, microtubules, or F- actin (Chikashige et al., 2006; Crisp et al., 2006; King et al., 2008; Morimoto et al.,

2012). Therefore, the SUN-KASH complexes are also known as linkers of the nucleoskeleton to the cytoskeleton (LINC).

The structure of the mammalian SUN-KASH complex was recently resolved (Sosa et al.,

2012; Zhou et al., 2012b). In this complex, three KASH domains are anchored in one cloverleaf-like trimer of SUN domains. The SUN domain protomer has several functional domains: an α-helical stalk, a compact β-sandwich core, a cation-loop, and a protruding anti-parallel β-sheet named the KASH-lid (Figure 1.2). The α-helix serves as an extension of the trimeric coiled-coil domain of SUN proteins, which facilitates the trimerization. The trimer is further stabilized by a large interacting surface on the β- sandwich core and the hydrogen bonds between the α-helix of one protomer to the β- sandwich of the adjacent protomer. The KASH domain is clamped between the KASH lid of one protomer and the β-sandwich core of the adjacent protomer. Without binding the

KASH domain, the KASH-lid conformation is rather random (Sosa et al., 2012). The very C-terminal “PPPX” motif is positioned in a KASH pocket formed by S641, Y703,

Y707, H628, and the cation-loop (C601-C705), which explains why these four amino acids are critical for SUN-KASH interactions (Sosa et al., 2012).

1.3 Function of opisthokont SUN and KASH proteins

1.3.1 Nuclear positioning mediated by opisthokont LINC complexes

Mammalian SUN1 and SUN2 interact with Lamin A, and the NE localization of SUN2 depends on Lamin A (Haque et al., 2006; Haque et al., 2010). SUN1 and SUN2 interact

4 with the mammalian KASH proteins Nesprin-1 and Nesprin-2 at the NE (Crisp et al.,

2006; Lei et al., 2009; Padmakumar et al., 2005). These two KASH proteins consist of N- terminal F-actin-binding calponin homology domains, a long stalk domain composed of spectrin repeats, and a C-terminal KASH domain (Crisp et al., 2006; Padmakumar et al.,

2005). The spectrin repeats each assemble into a three-helix bundle, which makes the protein flexible in length, perhaps involved in buffering mechanical stress (Autore et al.,

2013; Djinovic-Carugo et al., 2002; Grum et al., 1999; Lenne et al., 2000). Nesprin-1 and

-2 interact with F-actin and connect the INM lamins through SUN1 and SUN2 (Figure

1.1A) (Crisp et al., 2006; Zhang et al., 2007). These LINC complexes are responsible for anchoring the synaptic nuclei at the mouse neuromuscular junction (Lei et al., 2009;

Zhang et al., 2007). Nesprin-1 and -2 also link the centrosome to the nucleus through interactions with the dynein/dynactin complex (Figure 1.1A) (Zhang et al., 2009). This connection is essential to interkinetic nuclear migration and nucleokinesis in mice (Zhang et al., 2009). During fibroblast migration in response to wounds, the cell polarizes by reorienting the centrosome to the leading edge and repositioning the nucleus to the rear, which is facilitated by the transmembrane actin-associated nuclear (TAN) lines (Gomes et al., 2005). TAN lines are formed by SUN2 and Nesprin-2 at the NE and transfer the retrograde actin flow to the nucleus (Luxton et al., 2010).

In Drosophila, the localization of the KASH protein Klarsicht at the nuclear periphery depends on a type B lamin and a SUN protein, Klaroid (Patterson et al., 2004). Klaroid also forms nuclear aggregates in transgenic flies expressing a mutated Lamin C that lacks

5 the first 42 amino acids (Dialynas et al., 2010), suggesting that Klaroid might be associated with Lamin C. Klarsicht is connected to microtubules and is responsible for the apical nuclear migration during photoreceptor formation (Kracklauer et al., 2007;

Patterson et al., 2004). The nuclear periphery localization of another KASH protein,

MSP-300 also depends on Klaroid (Technau and Roth, 2008). MSP-300 is an ortholog of mammalian Nesprin-1 and -2. Although its role in nuclear anchorage was previously unclear (Technau and Roth, 2008; Xie and Fischer, 2008; Yu et al., 2006), a recent study revealed that MSP-300 interacts with D-Titin/Sallimus and anchors mitochondria and endoplasmic reticulum (ER) to the striated muscle Z-discs (Elhanany-Tamir et al., 2012).

It also cooperatively functions with Klarsicht to promote even nuclear spacing in striated muscle (Elhanany-Tamir et al., 2012).

In C. elegans, the SUN protein UNC-84 co-localizes with Ce-lamin at the NE and the localization of UNC-84 at the NE depends on Ce-lamin (Lee et al., 2002). UNC-84 recruits KASH proteins UNC-83 and ANC-1 to the NE (Figure 1.1A) (McGee et al.,

2006; Starr and Han, 2002). UNC-83 targets kinesin-1 and/or dynein to the nuclear periphery, and the force provided by the motor proteins drives the nuclear migration in C. elegans hypodermal P cells and embryonic hypodermal cells (Fridolfsson and Starr,

2010; Meyerzon et al., 2009; Starr et al., 2001). ANC-1 interacts with F-actin and is required for nuclear anchorage in the adult C. elegans syncytial hypodermis (Starr and

Han, 2002).

1.3.2 Chromosome movement mediated by opisthokont LINC complexes

SUN-KASH complexes can also link chromosomes to the cytoskeleton. In

Schizosaccharomyces pombe, centromeres are tethered to Sad1 through Csi1 (Figure

1.1B) (Hou et al., 2012). Loss of Sad1 or Csi1 leads to high-frequency centromere clustering defects (Hou et al., 2012). At meiotic prophase, telomeres are tethered to Sad1 through Bqt1, Bqt2, and telomere-associated proteins Taz1 and Rap1 (Chikashige et al.,

2006). Telomeres are further linked to dynein motors by KASH protein Kms1 and Kms2

(Figure 1.1B) (Ding et al., 1998; Miki et al., 2004; Shimanuki et al., 1997; Yamamoto et al., 1999). This results in telomere clustering and later nuclear oscillation between the cell poles, which facilitates homologous paring and recombination (Ding et al., 2004). In mammals, a recent study revealed a similar chromosomal bouquet formation mediated by a SUN1-KASH5 complex (Morimoto et al., 2012).

1.3.3 Nuclear pore complex assembly regulated by SUN1

In addition to nuclear positioning and chromosome movement, human SUN1 was found to be associated with the nuclear pore complexes (NPCs), and was required for the even distribution of NPCs (Liu et al., 2007). Coincident with this finding, depleting SUN1 led to impaired NPC assembly similar to the defects caused by depleting POM121, indicating that SUN1 plays a role in NPC assembly with POM121 during interphase (Rothballer et al., 2013; Talamas and Hetzer, 2011).

As a summary, Table 1.1 provides an overview of known SUN-KASH pairs from different organisms and their known or proposed function. 7

1.4 Plant SUN proteins and their functions

1.4.1 Plant SUN proteins

Two types of SUN protein have been identified in plant genomes: the canonical C- terminal SUN domain (CCSD) type and the plant-prevalent mid-SUN 3 transmembrane

(PM3) type (Graumann et al., 2010a; Murphy et al., 2010). The CCSD type has a SUN domain at the C-terminus, while the signature of the PM3 type is a centrally positioned

SUN domain followed by a highly conserved domain of unknown function (Murphy et al., 2010). The PM3 type was first discovered in maize, but is also present in other plant species, as well as in opisthokonts (Murphy et al., 2010). PM3-type SUN proteins have not yet been functionally investigated. To date, all well-studied SUN proteins belong to the CCSD type, and therefore, only plant CCSD-type SUN proteins will be discussed in detail here.

The Arabidopsis genome encodes two CCSD-type SUN proteins—AtSUN1 and AtSUN2

(Graumann et al., 2010a; Oda and Fukuda, 2011). They share a similar domain organization to non-plant SUN proteins: an N-terminal domain with a nuclear localization signal (NLS), a transmembrane domain, a coiled-coil domain, and a SUN domain. Deleting the N-terminal domain, the NLS, or the coiled-coil domain of AtSUN1 or AtSUN2 affect their NE localization, suggesting that these domains are involved in NE targeting and integration. Both genes are expressed ubiquitously in various tissues, including roots, hypocotyl, cotyledons, and leaves. As analyzed by fluorescence resonance energy transfer assays, AtSUN1 and AtSUN2 can form homo- or hetero-

8 complexes (Graumann et al., 2010a), perhaps similar to the mammalian SUN protein trimers.

This is supported by the predicted 3D structure of the AtSUN1 SUN domain derived from comparative modeling using the SUN domain of Homo sapiens SUN2 (HsSUN2) as template (Figure 1.2A). According to the HsSUN2 model, the SUN2 trimer harbors three binding sites for the KASH domain, such that the SUN-KASH complex is a hexamer

(Sosa et al., 2012). AtSUN1 likely harbors the same binding configuration, because the essential structures and amino acids involved in KASH binding are mostly conserved

(Figure 1.2B and Figure 1.3): (1) the KASH-binding pocket is very well-conserved; (2) the cation loop is conserved, especially within plant SUN domains; (3) although the residues of the KASH lid are not conversed, a corresponding fragment is present in all plant SUN domains (Figure 1.3). In contrast, the residues that are dispensable for SUN-

KASH interaction—residue C563 of HsSUN2 that forms a disulfide bond with a cysteine in the KASH domain and residue N636 of HsSUN2, the N-glycosylation site—are not conserved in plant SUN domains (Figure 1.3).

1.4.2 Possible function of plant SUN proteins in mitosis and meiosis

Mammalian SUN1 is associated with NPCs and interacts with Lamins (Liu et al., 2007).

It is suggested that this kind of INM protein links mitotic ER to chromatin in telophase and might mediate NE reassembly during mitosis (Guttinger et al., 2009). Plant SUN proteins may also be involved in a similar process. In Arabidopsis, AtSUN1 diffuses to the ER after NE breakdown and mainly localizes to the distal side of the separated

9 chromosomes throughout anaphase (Oda and Fukuda, 2011). At telophase, an enriched

AtSUN1 signal starts to enclose the chromosomes from the distal surface to the proximal surface. At the same time, the signal at the ER becomes gradually reduced, indicating the translocation of AtSUN1 to the newly formed NE (Oda and Fukuda, 2011). Studies of

AtSUN1 and AtSUN2 in BY-2 cells showed similar results (Evans and Graumann, 2011).

In addition, studies using BY-2 cells also showed that AtSUN1 and AtSUN2 were associated with membranes around the spindle and close to the chromosomes (Evans and

Graumann, 2011).

Similar to mammalian telomere organization, plant telomeres are also attached to the NE during meiosis (Bass, 2003; Bozza and Pawlowski, 2008). During maize (Zea mays L.) meiotic prophase I, starting at the end of leptotene, telomeres begin to cluster at the nuclear periphery to form a bouquet (Bass et al., 1997). This process persists through zygotene and ends at early pachytene when telomeres start to disperse at the nuclear periphery (Bass et al., 1997). In Arabidopsis, telomeres move to the nuclear periphery and form a loose cluster, perhaps a temporary bouquet (Roberts et al., 2009).

1.5 Nuclear positioning in plants

In the plant life cycle, a considerable number of developmental events require or are accompanied by nuclear positioning, including fertilization, root and leaf hair growth, and asymmetric divisions. Besides development, nuclear relocation is also stimulated by environmental factors such as pathogen invasion, symbiotic microbe interaction, and mechanical stimuli.

1.5.1 Nuclear movement of pollen tubes

The most obvious nuclear movement event is the long-distance traveling of the sperm cells (SCs) in pollen tubes, a process accompanied by the vegetative nucleus (VN). There are two types of angiosperm pollen—tricellular and bicellular. The tricellular pollen contains a VN and two SCs, while the bicellular pollen contains a VN and a generative cell (GC) which undergoes mitosis to generate two SCs during pollen tube growth

(McCue et al., 2011). Evidence from studies on various angiosperm species shows that the VN and GC/SCs are closely associated, and in many cases they are physically connected in both pollen grains and pollen tubes (McCue et al., 2011). Therefore, they were named the “male germ unit” (MGU) (Dumas et al., 1985) and the MGU does travel as a unit in pollen tubes. However, the traveling order of VN and GC/SCs seems to be species-specific. In growing pollen tubes of many angiosperm species, the VN precedes

(close to the growing pollen tube tip) the GC/SC (Heslop-Harrison and Heslop-Harrison,

1989b; McCue et al., 2011), but the order can be complicated in some species. During

Amaryllis vittata Ait. pollen tube growth, either the VN or the GC enters the tube first, but the VN always precedes the GC afterwards. When the GC undergoes mitosis, the position of VN and GC starts shifting. A similar process was also observed during tobacco (Nicotiana tabacum) pollen tube growth—the VN preceded the GC in the beginning and later became proximal to the dividing GC (Palevitz, 1993). The function of

VN repositioning during GC division is unknown.

The molecular mechanism underlying the MGU movement has been studied for decades, but still remains an enigma. Depolymerizing tobacco pollen microtubules using 0.5 µM oryzalin impaired VN and GC movement but did not affect pollen tube growth and microfilament organization (Astrom et al., 1995). In another study, 1 µM oryzalin treatment of tobacco pollen led to similar results but with presence of abnormal microfilaments (Kaul et al., 1987). During Galanthus nivalis L. pollen tube growth, the

VN was observed to precede the SC in most cases (Heslop-Harrison et al., 1988).

Depolymerizing microtubules using 0.5 mM colchicine affected this order and increased the distance between the leading nucleus and the growing pollen tube apex (Heslop-

Harrison et al., 1988). The VN and GC were more separated when treated with 1 mM colchicine (Heslop-Harrison et al., 1988). However, at these two colchicine concentrations the pollen tube growth was also affected (Heslop-Harrison et al., 1988).

Nevertheless, these studies suggest that a microtubule network is responsible for MGU movement. The identification of dynein-related polypeptides in tobacco was proposed to support this hypothesis, though no evidence was shown to link these polypeptides to nuclear movement (Moscatelli et al., 1996). On the contrary, anti-myosin antibodies labeled discrete foci at the VN periphery of tobacco pollen, suggesting that myosin and microfilament might be responsible for the VN movement (Tirlapur et al., 1995).

Supporting this hypothesis, immunoblot and immunolabeling using antibodies raised against non-plant myosins also identified putative myosins at periphery of the VN in

Lilium longiflorum and Helleborus foetidus L. (Heslop-Harrison and Heslop-Harrison,

1989c; Miller et al., 1995). Depolymerizing microfilaments of Lilium henryi Baker. using 12 cytochalasin D caused rapid contraction of the elongated VN, increased separation between the VN and GC, and reduced distance between the VN and the pollen tube tip

(Heslop-Harrison and Heslop-Harrison, 1989b). In addition, the Lotus japonicas symbiotic mutant, crinkle (crk) also has aberrant F-actin organization in pollen tubes and

GC movement defects (Ishikawa et al., 1993). In spite of all these studies, no specific genes have been identified in MGU movement, and whether the MGU movement is F- actin-dependent or microtubule-dependent is still unclear.

1.5.2 Nuclear movement of root and leaf hair

Another example of long-range nuclear movement is in root hairs. In Arabidopsis, the nucleus of a growing root hair is positioned at a relatively fixed distance from the apex, and interrupting this nuclear positioning by depolymerizing F-actin or trapping the nucleus with a laser beam prohibits root hair growth (Chytilova et al., 2000; Ketelaar et al., 2002). In Arabidopsis mature long root hairs, the nuclei still undergo dynamic movement (Chytilova et al., 2000), and in Medicago mature root hairs, the nucleus can be found at random positions (Sieberer and Emons, 2000) probably due to its dynamic movement property. Similarly, during the development of Arabidopsis trichomes— branched, single-cell leaf hairs—the nucleus migrates to a position close to the first branch point (Folkers et al., 1997).

A mechanism that regulates nuclear movement was recently identified. In Arabidopsis, myosin XI-I, which is recruited by WPP-domain interacting tail-anchored protein 1

(WIT1) and WIT2 to the NE, is responsible for the nuclear movement, at least in the

13 mature root zone, and mesophyll cell nuclear positioning in dark (Tamura et al., 2013).

Since, WIT1 interacts with WIP1, WIP2, and WIP3 (Zhao et al., 2008), myosin XI-I is proposed to be linked to the SUN-WIP NE bridges (CHAPTER 2) through WIT1 and

WIT2 (Figure 1.1C) (Tamura et al., 2013).

1.5.3 Nuclear positioning during asymmetric cell divisions

Asymmetric cell divisions, which require the nucleus to be positioned at the future division site, play an important role in plant development. During lateral root formation, the nuclei of two neighboring lateral root founder cells migrate toward the common cell boundary. This is followed by asymmetric cell divisions leading to two small, adjacent cells and two larger, peripheral cells (De Rybel et al., 2010; De Smet et al., 2007). During pollen mitosis I, the nucleus in microspores migrates to allow asymmetric division to produce a large vegetative cell and a small generative cell. Arabidopsis MICROTUBULE

ORGANIZATION 1 (MOR1; also known as GEM1) and its tobacco ortholog

TOBACCO MICROTUBULE BUNDLING POLYPEPTIDE OF 200 kDa (TMBP200) are required for this process, and mutations in these proteins lead to defects in pollen production (Oh et al., 2010; Park et al., 1998; Twell et al., 2002). Guard cell formation also involves several steps of asymmetric cell division (reviewed in refs. De Smet and

Beeckman, 2011; Scheres and Benfey, 1999). In monocots, the guard mother cells are yielded by one asymmetric division and then divide symmetrically to produce guard cell pairs. Subsequently, the adjacent epidermal cells divide asymmetrically to generate subsidiary cells. In dicots, a protodermal cell divides asymmetrically to produce a

14 meristemoid, which is capable of generating either epidermal pavement cells or guard mother cells. Molecular factors controlling nuclear movement in these processes are unknown.

1.5.4 Root nuclear movement responding to microbes

Nuclear movement is also actively involved in plant-microbe interactions. Well studied cases are Rhizobium-induced nodulation (Gage, 2004) and arbuscular mycorrhizal symbiosis (Smith et al., 2006). In both cases, the nuclei are known to guide the development of infection apparatus inside root cells.

Upon Rhizobium infection, growing root hairs curl and develop infection threads, intracellular tubules where rhizobia grow and divide (Gage, 2004). The root hair nuclei are then uncoupled from maintaining a distance with the growing tips (see 1.5.2), and instead they strictly guide the growth of infection threads into root cortex (Fåhraeus,

1957; Nutman, 1959). The connection between the nuclei and infection threads is likely microtubules as shown by a study in Medicago (Timmers et al., 1999), while the basipetal nuclear movement towards the root cortex is proposed to be F-actin-dependent

(Lloyd et al., 1987). Interestingly, uncoupled nuclear movement at the growing root hair tips was also observed during the infections of root-knot nematodes in Lotus japonicas, suggesting that this movement is not a process unique to symbiosis (Weerasinghe et al.,

2005).

During arbuscular mycorrhizal fungal infection, when fungi penetrate host root cells, the host cell nucleus rapidly positions itself beneath the appressorium contact site and 15 promotes cytoskeleton and ER rearrangements (Genre et al., 2008; Genre et al., 2005).

The nucleus then migrates away from the contact site, accompanied by a formation of a column comprised of cytoskeletal elements and ER (Genre et al., 2008; Genre et al.,

2005). This newly formed structure strictly defines the future intracellular path of hyphal penetration and is called the pre-penetration apparatus (Genre et al., 2008; Genre et al.,

2005). However, in case of a pathogenic fungus, the host cell nucleus stays at the appressorium contact site, which is accompanied by cell wall thickening and papilla formation that blocks fungal penetration (Schmelzer, 2002).

1.5.5 Nuclear positioning responding to physical stimuli

The position of plant organelles is light dependent, and the nucleus is not an exception.

Light-induced nuclear positioning was first discovered in the alga Vaucheria sessilis.

Point illumination of low intensity of blue light attracted the nucleus and other organelles to the illuminated area (Blatt and Briggs, 1980). Later, light dependent nuclear movement was found in the fern Adiantum capillus-veneris. The nuclei locate to the anticlinal walls under dark or strong light conditions and to the periclinal walls under weak light conditions (Kagawa and Wada, 1993; Kagawa and Wada, 1995; Tsuboi et al., 2007).

Photoreceptor nemochrome1 and phototropin2 were assigned to regulating the nuclear movement under different light conditions (Tsuboi et al., 2007). Arabidopsis has similar light-dependent nuclear positioning. In dark, the nuclei reside at the periclinal walls, while under strong blue-light the nuclei are repositioned to the anticlinal walls (Iwabuchi

16 et al., 2010; Iwabuchi et al., 2007). Blue-light induced nuclear movement is phototropin2 and F-actin dependent (Iwabuchi et al., 2010; Iwabuchi et al., 2007).

Other physical stimuli such as wounding and local pressure application at the cell surface cause nuclear migration to the “incident site”. The nuclei of the cells adjacent to the wounded site were reported to migrate towards the wounded site (Nagai, 1993). It has also been demonstrated in multiple plant species that applying local pressure at the cell surface using a needle led to nuclear migration to the contact site (Gus-Mayer et al.,

1998; Hardham et al., 2008; Kennard and Cleary, 1997; Sato et al., 1999). This stimulus is thought to mimic pathogen attack (Gus-Mayer et al., 1998; Hardham et al., 2008).

1.6 Plant KASH analogs involved nuclear positioning

Despite the importance of nuclear positioning and chromosome movement during the plant life cycle, little is known about the underlining molecular mechanisms. Based on opisthokont studies, the LINC complexes are widely involved in these processes.

However, plant genomes do not encode any homologs of opisthokont KASH proteins.

This dissertation research focused on identifying plant KASH protein analogs and studying their function in nuclear positioning.

In CHAPTER 2, the first KASH protein analog in plants, the WPP domain-interacting protein (WIP) family, was identified. WIP family members contain a cytoplasmic coiled- coil domain, a transmembrane domain, and a short PNS tail, which terminates in a conserved “VPT” motif. WIPs are plant-specific ONM proteins and Arabidopsis has three homologs—WIP1, WIP2, and WIP3. Analogous to opisthokont KASH proteins, 17

WIP1 interacts with the SUN domain of SUN proteins through its PNS tail. The PNS tail of WIPs shares little homology to the PNS tails of reported opisthokont KASH proteins.

Therefore, to distinguish it from the KASH domain, the plant SUN domain-interacting tail present in WIPs was termed the SIT domain, and the plant KASH analogs were then named as SIT proteins.

Unlike opisthokont LINC complexes, the SUN-WIP NE bridges, are involved in anchoring the RanGTPase activating protein 1 (RanGAP1) to the NE (Figure 1.1C and

Table 1.1) (Xu et al., 2007; Zhou et al., 2012a). This mechanism of RanGAP anchoring is not evolutionary conserved (Hopper et al., 1990; Matunis et al., 1998; Rose and Meier,

2001). Thus, the first known function for plant NE bridging complexes already suggests that they have not only diverged in terms of the ONM SUN-binding partners, but also in terms of their cellular functions.

In CHAPTER 3, the nuclear movement function of WIPs was addressed in pollen tube growth. In pollen tubes the sperm cells are associated with the vegetative nucleus and they move together as a unit. In this study, WIP1, 2, 3, and their ONM binding partners,

WPP domain-interacting tail-anchored protein 1(WIT1) and WIT2 were shown to mediate VN movement (Figure 1.1C and Table 1.1). Loss of WIP1, 2, 3 or WIT1, 2 led to impaired vegetative nuclear movement, which further caused inefficient sperm cell delivery.

In CHAPTER 4, based on the conserved characteristics of WIPs—a transmembrane domain and a conserved “VPT” motif, a computer program was developed to mine the

18 plant proteome for more plant SIT proteins. Ten new potential plant SIT proteins were identified, and four of them (NEMO, NEMOL, NEMOS, NEME) were experimentally verified (Figure 1.1C and Table 1.1). One SIT protein, NEMO1, was shown to be associated with F-actin filaments and play a role in guard cell nuclear anchorage (Figure

1.1C and Table 1.1).

These results revealed that plants incorporate a different set of SUN domain-interacting proteins at the ONM to accommodate functions similar to, but not restricted to, opisthokont KASH proteins. However, compared with what is still unknown, the knowledge obtained in this dissertation research is merely the beginning. The SIT proteins identified and functionally characterized here do provide a good start, and the perspectives are addressed in CHAPTER 5.

Figure 1.1 SUN-KASH/SIT bridges across the NE in different organisms. Since SUN domains are relatively conserved across species, all SUN proteins are drawn as trimers according to the evidence provided for HsSUN2 (Sosa et al., 2012; Zhou et al., 2012b). For simplicity, all KASH/SIT proteins are drawn as monomers. See text for details.

Figure 1.2 Computed three-dimensional model of the SUN domain of AtSUN1. (A) The S251- D453 fragment of AtSUN1 was modeled using MODELLER (Sali and Blundell, 1993). The SUN domain of HsSUN2 was used as a template (PDB: 4FI9). Three models were computed and the one with the lowest zDOPE score is shown. Magenta, model of the AtSUN1 SUN domain. Cyan, SUN domain of HsSUN2. Gray, Nesprin-2 KASH domain in the HsSUN2-KASH complex. (B) Computed surface of the binding pocket for the SIT C-terminus in AtSUN1. Red, V301-N318 fragment of AtSUN1 corresponding to the KASH lid of HsSUN2 (Y567-S587). Orange, S324-C333 fragment of AtSUN1, corresponding to the cation-loop of HsSUN2 (C601-C705). Purple, residue H360, S371, H439, and Y443 of AtSUN1, corresponding to H628, S641, Y703, Y707 of HsSUN2, respectively. Images were generated using UCSF Chimera package (Pettersen et al., 2004) and POV-Ray (http://www.povray.org/).

Figure 1.3 Amino acid sequence alignment of plant SUN domains with the SUN domain of HsSUN2.Alignment was performed using ClustalW2 (http://www.ebi.ac.uk/Tools/msa/clustalw2/) with default settings, except that the output sequences were kept as input order. Image was generated using JalView (Waterhouse et al., 2009) and ClustalX color. GI, NCBI GenInfo identifier. Black frames in the alignment are numbered at the top. Frame 1 indicates C563 in HsSUN2, which forms a disulfide bond with Homo sapiens Nesprin-2 C6862. This disulfide bond is dispensable for SUN-KASH interaction, and in plant SUN domains this position is instead a conserved D or E. Frame 2 indicates the KASH-lid in HsSUN2, however, the sequences have low similarity between HsSUN2 and plant SUN proteins. Frame 3 represents the cation loop in HsSUN2. This cation loop and residues indicated by frame 4, 6, 7, 8 (correspond to H628, S641, Y703, and Y707 of HsSUN2, respectively) form the pocket holding the KASH C-terminus and are well conserved in plant SUN domains. Frame 5 represents N636 of HsSUN2, the N-glycosylation site. N-glycosylation of HsSUN2 is dispensable for KASH binding, and this position is a conserved D in plant SUN domains.

Figure 1.3 Amino acid sequence alignment of plant SUN domains with the SUN domain of HsSUN2.

SUN KASH/SIT Cytoplasmic Partner: Function (Reference)

Mammal

SUN1/2 Nesprin-1 F-actin: anchoring the synaptic nuclei under the mouse neuromuscular junction (Zhang et al., 2007). Dynein/dynactin complex: connecting the nucleus to centrosome for to interkinetic nuclear migration and nucleokinesis (Zhang et al., 2009).

SUN1/2 Nesprin-2 F-actin: anchoring the synaptic nuclei under the mouse neuromuscular junction (Zhang et al., 2007). Dynein/dynactin complex and Kinesin: connecting the nucleus to centrosome for to interkinetic nuclear migration and nucleokinesis (Zhang et al., 2009).

SUN1/2 Nesprin-3 Plectin, BPAG1, or MACF: connecting the nucleus to intermediate filaments or microtubules which stables the anchorage of the nucleus and maintains the structure/shape of the nucleus (reviewed in (Ketema and Sonnenberg, 2011)).

Maybe SUN1/2 Nesprin-4 Kinesin-1: predicted to promote nuclear migration toward the base of the secretory epithelial cells (Roux et al., 2009).

SUN1/2 KASH5 Dynein/dynactin complex: telomere movement during meiosis (Morimoto et al., 2012).

SUN3 Nesprin-1 Proposed to be kinesin II, Dynein/dynactin, or F-actin: probably links the nucleus to posterior manchette durng sperm head formation (Gob et al., 2010).

SUN1η Nesprin-3 Proposed to be plectin: proposed to be a non-NE complex anchoring acrosome to the anterior actin filaments during sperm head formation (Gob et al., 2010).

SPAG4 Unknown Unknown: Testis-specific, non-NE localized, function unknown (Shao et al., 1999).

continued

Table 1.1 Known SUN-KASH/SIT pairs from different organisms, their cytoplasmic partners, and their known or proposed functions.

Table 1.1 continued

SPAG4L/4L-2 Unknown Unknown: restricted to the apical nuclear region of round spermatids facing the acrosomic vesicle, and probably involved in linkage of the acrosomic vesicle to the spermatid nucleus and in acrosome biogenesis (Frohnert et al., 2011).

Drosophila

Klaroid MSP-300 F-actin: nuclear anchoring during Drosophila oogenesis (Yu et al., 2006).

Unknown MSP-300 D-Titin/Sallimus: anchoring mitochondria and endoplasmic reticulum to the striated muscle Z-discs (Elhanany-Tamir et al., 2012). Unknown: anchoring microtubules to the NE in striated muscle (Elhanany-Tamir et al., 2012). Unknown: Anchoring the nuclei to myofibril compartment in striated muscle (Elhanany-Tamir et al., 2012).

Klaroid Klarsicht Proposed to be microtubule motors: nuclear migration during eye development (Kracklauer et al., 2007; Patterson et al., 2004). Unknown: nonrandom sister chromatid segregation (Yadlapalli and Yamashita, 2013).

Unknown Klarsicht Unknown: anchoring microtubules to the NE in striated muscle (Elhanany-Tamir et al., 2012). Unknown: promoting even myonuclear spacing in both striated muscle and nonstriated myotubes (Elhanany-Tamir et al., 2012).

SPAG4/Giacomo Unknown Yuri and dynein-dynactin: involved in centriolar-nuclear attachment during spermatogenesis (Kracklauer et al., 2010).

C. elegans

UNC-84 ANC-1 F-actin: nuclear anchorage in the adult C. elegans syncytial hypodermis (Starr and Han, 2002).

UNC-84 UNC-83 Kinesin-1 and dynein: nuclear migration in embryonic C. elegans hypodermal cells (Meyerzon et al., 2009; Starr et al., 2001).

continued

Table 1.1 continued

SUN-1/matefin ZYG-12 Dynein and ZYG-12A: linkage between the centrosome and nucleus (Malone et al., 2003), meiotic chromosome paring and synapsis (Penkner et al., 2009; Sato et al., 2009), and nuclear positioning within the syncytial gonad (Zhou et al., 2009).

SUN-1/matefin KDP-1 Unknown: cell-cycle progression (McGee et al., 2009).

S. pombe

Sad1 Kms1 and Dynein and centrosomes: meiotic chromosome paring and Kms2 synapsis (Chikashige et al., 2006; Ding et al., 1998; Miki et al., 2004; Shimanuki et al., 1997; Yamamoto et al., 1999)

S. cerevisiae

Mps3 Unknown Unknown: Mps3 is involved in spindle pole body insertion into the NE and NE homeostasis (Friederichs et al., 2011); it interacts with Mps2 to connect the spindle pole body to the NE and function in spindle body duplication (Jaspersen et al., 2006).

Mps3 Csm4 Probably F-actin: meiotic telomeres are tethered to Mps3 at the NE by Ndj1 and further connected to cytoskeletons (perhaps actins) by Csm4 (Conrad et al., 2008) (Koszul et al., 2008; Wanat et al., 2008).

Mps3 Unknown Unknown: Mitotic telomeres are tethered to Mps3 at the NE by Sir4 and the telomere clustering is mediated by two Mps3 associated proteins (Bupp et al., 2007), Ebp2 and Rrs1 (Horigome et al., 2011).

Dictyostelium

SUN-1 Unknown Unknown: SUN-1 connects the centrosome to chromatin and ensures genome stability (Xiong et al., 2008)

Unknown Interaptin F-actin: Function unknown (Rivero et al., 1998)

Arabidopsis

continued

Table 1.1 continued

AtSUN1/2 AtWIP1/2/3 RanGAP: anchoring RanGAP to the NE (Zhou et al., 2012a) (see also Chapter 2).

AtSUN1/2 AtWIP1/2/3 AtWIT1 and AtWIT2: recruit myosin XI-I to the NE which regulates the root nuclear shape (Tamura et al., 2013) (see also CHAPTER 2) and nuclear movement in mesophyll cells after dark and in roots (Tamura et al., 2013)

AtSUN1/2 AtNEMO1 Associated with F-actin: anchor the nuclei to the center in guard cells (CHAPTER 4).

AtSUN1/2 AtNEMO2 Unknown: Unknown (see CHAPTER 4)

AtSUN1/2 AtNEMOL Unknown: Unknown (see CHAPTER 4)

AtSUN1/2 AtNEMOS Unknown: Unknown (see CHAPTER 4)

CHAPTER 2

Novel plant nuclear envelope bridges are involved in RanGAP anchoring and

nuclear shape determination

2.1 Abstract

Inner nuclear membrane Sad1/UNC-84 (SUN) proteins interact with outer nuclear membrane (ONM) Klarsicht/ANC-1/Syne-1 homology (KASH) proteins, forming linkers of the nucleoskeleton and the cytoskeleton which are conserved from yeast to humans and involved in the positioning of nuclei and chromosomes. Defects in SUN-KASH bridges are linked to muscular dystrophy, progeria, and cancer. SUN proteins were recently identified in plants, but their ONM KASH partners are unknown. Arabidopsis

WPP-domain interacting proteins (AtWIPs) are plant-specific ONM proteins that redundantly anchor Arabidopsis Ran GTPase activating protein 1 (AtRanGAP1) to the nuclear envelope (NE). Here, I report that AtWIPs are plant-specific KASH protein analogs interacting with Arabidopsis SUN proteins (AtSUNs). The interaction is required for both AtWIP1 and AtRanGAP1 NE localization. AtWIPs and AtSUNs are necessary for maintaining the elongated nuclear shape of Arabidopsis epidermal cells. Together, our data identify the first KASH protein analogs in the plant kingdom and provide a novel function of SUN-dependent NE bridging complexes, suggesting that a functionally diverged form of this complex is conserved beyond the opisthokonts.

2.2 Introduction

The nuclear envelope (NE) consists of an outer nuclear membrane (ONM) and an inner nuclear membrane (INM) that enclose a lumen called the periplasmic or perinuclear space (PNS) (Gerace and Burke, 1988). In non-plant eukaryotes, the ONM and INM are bridged by interactions between KASH (Klarsicht, ANC-1, and Syne/Nesprin homology) proteins and SUN (Sad1 and UNC-84) proteins (reviewed by Graumann et al., 2010b;

Razafsky and Hodzic, 2009; Starr and Fridolfsson, 2010). KASH proteins are integral membrane proteins of the ONM with a short C-terminal tail domain in the PNS. SUN proteins are INM proteins that contain at least one transmembrane domain (TMD), which positions their conserved C-terminal SUN domain inside the PNS. The interaction of the

PNS tail of KASH proteins with the SUN domains of SUN proteins stably associates

KASH proteins with the ONM and prevents their diffusion to the ER (Crisp et al., 2006;

McGee et al., 2006; Padmakumar et al., 2005).

Many SUN proteins interact with the nuclear lamins in the nucleoplasm, while KASH proteins interact with the cytoskeleton or cytoskeleton-associated proteins. The SUN-

KASH interactions are thus part of linker of nucleoskeleton to cytoskeleton complexes conserved from yeast to humans, functioning in nuclear positioning and chromosome movement (Crisp et al., 2006). The founding members of SUN-KASH protein pairs were identified in Caenorhabditis elegans. Interaction of the SUN protein UNC-84 with the actin-binding KASH protein ANC-1 is involved in nuclear anchorage. UNC-84 also interacts with the KASH protein UNC-83, which recruits kinesin-1 to transfer forces for

30 nuclear migration (Horvitz and Sulston, 1980; Malone et al., 1999; Sulston and Horvitz,

1981). Similarily, the Schizosaccharomyces pombe SUN-KASH bridges, formed by Sad1 and Kms, transfer dynein motor forces to telomeres for positioning telomeres to the spindle pole body (Chikashige et al., 2006; Miki et al., 2004).

SUN proteins were recently identified in plants (Graumann et al., 2010a; Moriguchi et al., 2005; Murphy et al., 2010). The two Arabidopsis SUN proteins—AtSUN1 and

AtSUN2—share the protein structure of the non-plant SUN proteins: an N-terminal domain containing a NLS, a TMD, a coiled-coil domain, and a conserved SUN domain.

Although both SUN proteins are expressed in almost all Arabidopsis tissues (Graumann et al., 2010a), a reported AtSUN1/AtSUN2 double mutant shows no phenotypes except for a nuclear shape change in root hairs which does not affect root hair growth (Oda and

Fukuda, 2011). No plant KASH proteins were identified. Therefore, the function of plant

SUN proteins is unclear.

Arabidopsis WPP-domain interacting proteins (AtWIPs) are three plant-specific ONM proteins that redundantly anchor Arabidopsis Ran GTPase activating protein 1

(AtRanGAP1) to the NE (Xu et al., 2007). We show here that AtWIP1, AtWIP2, and

AtWIP3 interact with AtSUN1 and AtSUN2 at the NE. The interaction is mediated through the highly conserved, yet plant-specific, PNS tail of AtWIPs and an extended

SUN domain of AtSUNs, which is conserved among land plants. The AtSUN-AtWIP1 interaction is required for the NE localization of AtWIP1 and consequently the NE localization of AtRanGAP1, a function for SUN-dependent NE bridges not reported in

31 any other organism. Mimicking a sun1-KO sun2-KD mutant, loss of nuclear polarity was also found in a triple loss-of-function wip1-1 wip2-1 wip3-1 mutant, indicating that

AtSUNs and AtWIPs are responsible for maintaining the elongated shape of plant nuclei.

Together, our data identified AtWIPs as the first plant KASH protein analogs and present novel functions of the SUN-dependent NE bridging complexes, suggesting that SUN- dependent NE bridges are conserved beyond the opisthokonts, but have functionally diverged.

2.3 Material and Methods

2.3.1 Plant materials

Arabidopsis (Columbia ecotype) were grown at 25⁰C in soil under 16-h light and 8-h dark or on MS (Caisson laboratories) plates under constant light. Mutant wip1-1 wip2-1 wip3-1 was reported previously (Xu et al., 2007). The sun1-KO sun2-KD mutant

(SALK_123093 for sun1-KO and SALK_049398 for sun2-KD) was a gift from Dr. Susan

Armstrong (University of Birmingham, UK).

2.3.2 Constructs

GFP-AtWIP1, GFP-TDFAtWIP1, and GFP-AtWIP1VVPT were previously described (Xu et al., 2007). CFP-AtSUN2, CFP-AtSUN2N, CFP-AtSUN2CC, CFP-

AtSUN2CSUN, AtSUN1-YFP, and AtSUN2-YFP were described by Graumann et al.

(2010a). WPP-GFP was previously reported as AtRanGAP1C-GFP (Rose and Meier,

2001). PCR was used to amplify AtRanGAP1, AtWIP1XT, Flag-AtSUN1, Flag-

AtSUN1NSUN, Myc-AtSUN2, Myc-AtSUN2NSUN, and NLS-GFP. All PCR products were cloned into the pENTR/D-TOPO vector (Invitrogen), confirmed by sequencing, and moved to destination vectors described by Karimi et al. (2002) and

Nakagawa et al. (2007) by LR reaction (Invitrogen): AtRanGAP1 was cloned into the pK7FWG2 vector to obtain AtRanGAP1-GFP; AtWIP1XT and NLS-GFP were cloned into the pK7WGF2 vector to obtain GFP-AtWIP1XT and GFP-NLS-GFP; Flag-AtSUN1,

Flag-AtSUN1NSUN, Myc-AtSUN2, and Myc-AtSUN2NSUN were cloned into the pK7WGR2 vector to obtain RFP-Flag-AtSUN1, RFP-Flag-AtSUN1NSUN, RFP-Myc-

AtSUN2, and RFP-MycAtSUN2NSUN; Flag-AtSUN1 was cloned into pGWB21 to obtain Myc-Flag-AtSUN1. AtWIP1 and AtWIP1XT were cloned to the pH2GW7 vector to obtain the overexpressing constructs.

2.3.3 Generation of transgenic plants

GFP-AtWIP1 and GFP-AtWIP1VVPT transgenic wild-type Arabidopsis, and

AtRanGAP1-GFP transgenic wip1-1 wip2-1 wip3-1 were previously described by Xu et al. (2007). AtRanGAP1-GFP transgenic wild-type Arabidopsis was previously described by Jeong et al. (2005). Other transgenic Arabidopsis were obtained by Agrobacterium- mediated floral dip (Clough and Bent, 1998).

2.3.4 Co-immunoprecipitation experiments

Agrobacterium cultures containing plasmids expressing the proteins of interest were co- infiltrated transiently into N. benthamiana leaves as described previously (Sparkes et al.,

2006). Plants were grown 3 days after infiltration, and leaves were collected and ground in liquid nitrogen into powders and Co-IP was performed at 4⁰C. For samples involving

Myc-Flag-AtSUN1, radioimmunoprecipitation assay buffer containing 50 mM Tris-HCl, pH7.5, 150 mM NaCl, 0.1% SDS, 0.5% NaDeoxycholate, 1% NP-40, 1 mM PMSF, and

1% protease inhibitor cocktail (Sigma-Aldrich) was used. Other protein extracts were prepared in NP-40 buffer containing 50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.5% NP-

40, 1 mM EDTA, 3 mM DTT, 1 mM PMSF, and 1% protease inhibitor cocktail (Sigma-

Aldrich). One-tenth of the protein extracts was used as input sample, and the rest were used for IP. Protein complexes were IP from extracts by anti-GFP antibody (ab290,

Abcam Cambridge) bound to A-sepharose beads (GE Healthcare). The immunoprecipitates were then analyzed by western blot with anti-AtWIP1 (1:2000, previously described by Xu et al. (2007)), anti-GFP (1:2000, 632569, Clontech), anti-

Myc (1:1000, M5546, Sigma-Aldrich), or anti-FLAG (1:2000, F7425, Sigma-Aldrich) antibody.

2.3.5 DAPI staining and nuclear length measurement

For leaf nuclear staining, fully expanded leaves before bolting were cut into small pieces.

For root hair nuclear DAPI staining, 7- or 8-day old Arabidopsis seedlings grown on MS plates were used. All samples were stained in 1 µg/ml DAPI solution for 20 min. Images were taken by a Nikon DS-Qi1Mc digital camera. The length of the nuclei was measured using Nikon NIS-Elements software.

2.3.6 Root hair and trichome nuclear positioning Assay

Young root hairs of 11-day old Arabidopsis seedlings carrying the GFP-NLS-GFP marker were imaged using Nikon Eclipse C90i confocal microscope. For trichome nuclear positioning assay, the nuclei of fully expanded young leaves of 25-day old plants were imaged using a Nikon DS-Qi1Mc digital camera, and distances were measured using Nikon NIS-Elements software.

2.3.7 Confocal microscopy and FRAP

Nikon Eclipse C90i confocal microscope with minimum or medium aperture was used to image 7- or 8-day old Arabidopsis seedlings. Intensities were measured using Nikon NIS- elements software. Cytoplasmic intensities shown in Figure 2.4 (the C1, C2 points) were measured close to the cell wall to capture maximal cytoplasmic values away from more central vacuoles. To reduce noise, intensity profiles shown in Figure 2.4 were calculated by averaging the intensities of adjacent 4 pixels. FRAP imaging was performed as described by Graumann et al. (2007).

2.4 Results

2.4.1 Identification of AtWIPs as ONM AtSUN-interacting partners

In most animal KASH proteins, the PNS tail terminates with a “PPPX” motif that is essential for SUN protein interaction and is required for NE localization of KASH proteins (Razafsky and Hodzic, 2009; Starr and Fridolfsson, 2010). Plant WIP family proteins are the only currently known plant ONM proteins that have a C-terminal PNS

35 tail (Xu et al., 2007). Although the tail is much shorter than those of animal KASH proteins (~9 vs. ~30 amino acids), they do terminate in four highly conserved amino acids—“VPT” ( representing any hydrophobic amino acid, and deletion of these four amino acids of AtWIP1 has been shown to diminish its NE localization (Xu et al., 2007).

As shown in Figure 2.1, the AtWIP1 PNS tail shows a low degree of similarity to known

KASH domains. The PNS tail is significantly shorter than the tail of most KASH proteins, but has similar length to that of C. elegans ZYG-12B, and KDP-1 and Interaptin from Dictyostelium discoideum (McGee et al., 2009; Minn et al., 2009; Xiong et al.,

2008). Specifically, the penultimate proline is highly conserved and a terminal serine/threonine residue is often present.

To determine whether AtWIPs bind AtSUNs, we tested the interaction between AtWIP1 and AtSUN2 by co-immunoprecipitation (Co-IP). As a control, a GFP-fused AtWIP1 mutation—GFP-AtWIP1XT—was made. In GFP-AtWIP1XT (Figure 2.2A), the PNS tail of AtWIP1 (PEPDTVVPT) was replaced by the ER luminal tail (RFYTKSAEA) of the tail-anchored ER protein Cytochrome b5c from Aleurites fordii (Hwang et al., 2004). To investigate the importance of the conserved last four amino acids, GFP-AtWIP1VVPT

(AtWIP1 without C-terminal “VVPT”, Figure 2.2A) was tested. AtSUN2 was C- terminally fused to a red fluorescence protein (RFP)-Myc tag (RFP-Myc-AtSUN2) and then transiently co-expressed in Nicotiana benthamiana with GFP-AtWIP1, GFP-

AtWIP1XT, and GFP-AtWIP1VVPT, respectively. After IP with anti-GFP antibody, co-immunoprecipitated AtSUN2 was detected by anti-Myc antibody. GFP-AtWIP1

36 bound AtSUN2, while GFP-AtWIP1XT did not bind (Figure 2.2C). Deletion of the conserved “VVPT” greatly diminished the interaction (Figure 2.2C). Thus, AtWIP1 interacts with AtSUN2, and the PNS tail of AtWIP1 is essential for the interaction.

Deleting “VVPT” only partially affected the interaction, indicating that the less conserved segment between “VVPT” and the TMD is also involved in binding AtSUN2.

Next, GFP-AtWIP2 and GFP-AtWIP3 were tested in co-IP assays with RFP-Myc-

AtSUN2 alongside GFP-AtWIP1 and GFP-AtWIP1XT (Figure 2.2D). GFP-tagged

AtWIP1, AtWIP2, and AtWIP3 co-immunoprecipitated AtSUN2, while GFP-AtWIP1XT did not, indicating that all three AtWIP family members bind AtSUN2. To determine whether AtWIPs also interact with AtSUN1, a Myc-Flag-tagged AtSUN1 (Myc-Flag-

AtSUN1) was co-immunoprecipitated with GFP-AtWIP1, GFP-AtWIP1XT, and GFP-

AtWIP1VVPT (Figure 2.2E). After IP by the anti-GFP antibody, AtSUN1 was detected using an anti-Myc antibody. Figure 2.2E shows that both the exchange of the PNS tail and the deletion of the “VVPT” motif greatly reduced binding between AtWIP1 and

AtSUN1. Figure 2.2F shows that GFP-AtWIP1, GFP-AtWIP2, and GFP-WIP3 all interact with RFP-Flag-AtSUN1. We conclude that all three AtWIPs are binding partners of both AtSUNs and that the PNS tail of AtWIP1 is important for binding.

2.4.2 AtSUNs interact with AtWIP1 through their SUN domain on the NE

Based on SUN-KASH interaction studies, the SUN domain or the segment between the coiled-coil domain and the SUN domain are required for interacting with KASH proteins

(Padmakumar et al., 2005; Stewart-Hutchinson et al., 2008). Protein sequence alignment

37 shows that unlike animal and fungal SUN proteins, sequence conservation among plant

SUN proteins begins immediately after the predicted coiled-coil domain (Figure 2.6). To test whether this extended plant SUN domain is required for binding WIPs, deletions of its N-terminal part (P266-R309, denoted as AtSUN2NSUN) and its C-terminal part

(deletion of R310-A455, denoted as AtSUN2CSUN) were constructed in AtSUN2

(Figure 2.2B and Figure 2.6). CFP-AtSUN2, CFP-AtSUN2N (deletion of the domain N- terminal to the TMD, Figure 2.2B), CFP-AtSUN2CC (deletion of the coiled-coil domain, Figure 2.2B), GFP-AtSUN2NSUN, and CFP-AtSUN2CSUN were tested for co-IP with AtWIP1. AtSUN2, AtSUN2N, and AtSUN2CC, but not GFP-

AtSUN2NSUN and CFP-AtSUN2CSUN bound AtWIP1 (Figure 2.2G). Hence, the

SUN domain is essential for interacting with AtWIP1.

2.4.3 AtSUN1 affects the mobility of AtWIP1 at the plant NE (data generated by

Katja Graumann)

The mobility of a membrane protein will be reduced upon interacting with other proteins

(Reits and Neefjes, 2001). To confirm that the SUN-WIP interaction occurs at the NE, we measured the mobility of AtWIP1-based fusion proteins using FRAP in the absence and presence of AtSUN-based fusion proteins. Firstly, we examined the mobility of GFP-

AtWIP1, GFP-AtWIP1VVPT, and GFP-TDFAtWIP1 (TDF, transmembrane-domain fragment, is defined as the TMD plus PNS tail, as shown in Figure 2.1 and Figure 2.2A) expressed transiently in N. benthamiana leaves and found that full-length GFP-AtWIP1 is

38 the least mobile (Figure 2.3A). (To quantify the mobility change, the maximum recovery was compared.) GFP-AtWIP1VVPT is significantly more mobile than GFP-AtWIP1 (t- test, p<0.01, n=30), due to the possibility that this deletion disrupts the interaction between AtWIP1 and N. benthamiana SUN proteins. Interestingly, GFP-TDFAtWIP1 is also more mobile (t-test, p<0.01, n=30) indicating that the cytoplasmic N-terminus of

AtWIP1 is involved in strong binding interactions. These have previously been shown to include RanGAP and WPP-Interacting tail-anchored protein (Xu et al., 2007; Zhao et al.,

2008).

Next, the GFP-AtWIP1-based constructs were co-expressed with RFP-Flag-AtSUN1 to investigate if mobility is affected (see Figure 2.7A for protein localization). A significant decrease (t-test, p<0.01, n=30) in mobility was detected for GFP-AtWIP1 upon co- expression with RFP-Flag-AtSUN1 (Figure 2.3B) indicating that AtSUN1 interacts with

GFP-AtWIP1 at the NE. To examine the effect of the SUN domain on GFP-AtWIP1 mobility at the NE, we co-expressed GFP-AtWIP1 with RFP-Flag-AtSUN1NSUN (see

Figure 2.7A for protein localizations). As shown in Figure 2.3B, GFP-AtWIP1 is more mobile when co-expressed with RFP-Flag-AtSUN1NSUN than with full length RFP-

Flag-AtSUN1 (t-test, p<0.01, n=30) and has the same mobility as when expressed on its own (t-test, p>0.05, n=30). The same effects were observed when expressing GFP-

AtWIP1 with either RFP-Myc-AtSUN2 (t-test, p<0.01, for GFP-AtWIP1 single expression, n=60; for co-expression with RFP-Myc-AtSUN2, n=30), or with RFP-Myc-

AtSUN2NSUN (t-test, p>0.05, for GFP-AtWIP1 single expression, n=60; for co-

39 expression with RFP-Myc-AtSUN2NSUN, n=30) as shown in Figure 2.3C. When GFP-

AtWIP1VVPT was co-expressed with either RFP-Flag-AtSUN1 or RFP-Myc-AtSUN2 the mobility of GFP-AtWIP1VVPT did not change (Figure 2.3D, t-test, P>0.05, n=30) indicating that the mobility change based on co-expressing the SUN proteins requires the

C-terminal VVPT motif. The mobility of the highly mobile GFP-TDFAtWIP1 also decreased when co-expressed with either RFP-Flag-AtSUN1 or GFP-Myc-AtSUN2

(Figure 2.3E, t-test, p<0.01, n=30), Together, these data corroborate the co-IP interaction results and show that the both the NSUN domain and the VVPT motif are required for the interaction of AtWIP1 with AtSUN1 and AtSUN2 at the plant NE.

2.4.4 AtSUN1 and AtSUN2 are required to anchor AtWIP1 to the NE

Previous studies have shown that the NE localization of KASH proteins is dependent on

SUN proteins (Crisp et al., 2006; Ketema et al., 2007; Padmakumar et al., 2005; Stewart-

Hutchinson et al., 2008). In GFP-AtWIP1VVPT transformed wild-type Arabidopsis, the

GFP signal at the NE is significantly reduced compared to GFP-AtWIP1 and diffuse signal appears in the cytoplasm, consistent with the VVPT motif being important for

AtWIP1 NE localization (Xu et al., 2007). To test if the NE localization of AtWIP1 depends on SUN proteins, we transformed GFP-AtWIP1 into a sun1-KO sun2-KD mutant. The mutant contains two T-DNA insertions which cause the complete absence of

AtSUN1 transcript and a reduction in the amount of AtSUN2 transcript (Figure 2.7B).

Three GFP-AtWIP1-expressing lines were obtained, and the GFP-AtWIP1 signal was imaged in undifferentiated root tip cells. Compared to GFP-AtWIP1-transformed wild

40 type seedlings, the GFP-AtWIP1 signal in the sun1-KO sun2-KD mutant is predominantly diffuse in the cytoplasm, very similar to the localization of GFP-

AtWIP1VVPT in wild type (Figure 2.4A). To quantify the NE concentration of the

GFP-AtWIP1 signal, we defined an NE localization index (NLI): the sum of the maximum from two NE intensities divided by the sum of the maximum cytoplasmic intensities ([N1+N2]/[C1+C2] as indicated in the wild-type intensity profile panel in Figure

2.4A). The more the signal is concentrated at the NE, the higher the NLI will be, with no apparent concentration having an NLI value close to 1. Figure 2.4 A and B show that the

NLI is significantly higher in GFP-AtWIP1-transformed wild type than in GFP-AtWIP1- transformed sun1-KO sun2-KD mutant and GFP-AtWIP1VVPT-transformed wild type

(t-test, P<0.01, n=50). The difference between GFP-AtWIP1-transformed sun1-KO sun2-

KD and GFP-AtWIP1VVPT-transformed wild type likely reflects the fact that sun2-KD is not a null allele. This result indicates that AtSUNs are required for the concentration of

AtWIP1 at the NE.

2.4.5 AtSUNs are required for AtRanGAP1 NE localization

In both the plant and animal lineages, RanGAP is associated with the ONM, proposed to be important for efficient RanGTP hydrolysis during nucleocytoplasmic transport (Hutten et al., 2008; Mahajan et al., 1997). Different from animals whose RanGAP is anchored by

RanBP2 at the NE, AtWIPs are required for anchoring AtRanGAP1 and AtRanGAP2 at the NE in undifferentiated root cells in Arabidopsis (Meier et al., 2010; Xu et al., 2007).

The loss of AtWIP1 at the NE in sun1-KO sun2-KD suggests that in plants, SUN proteins

41 may play a role in RanGAP NE localization. Hence, we examined the GFP signal in undifferentiated root cells of AtRanGAP1-GFP-transformed sun1-KO sun2-KD lines (10 independent lines were examined) and compared it with AtRanGAP1-GFP-transformed wild type and wip1-1 wip2-1 wip3-1. The NLI was used to quantitatively compare the signals. As shown in Figure 2.4 C and D, the AtRanGAP1-GFP signal was significantly more diffuse in the cytoplasm in both mutants than in wild type (t-test, P<0.01, n=55).

Again, the higher NLI of sun1-KO sun2-KD lines than that of wip1-1 wip2-1 wip3-1 lines may be due to that sun2-KD is not a null allele.

AtWIP1 interacts with the WPP domain of AtRanGAP1 through its N-terminal cytoplasmic coiled-coil domain (Xu et al., 2007), while AtSUN2 binds the PNS tail of

AtWIP1. We therefore tested whether AtSUN2 interacts with AtRanGAP1 through

AtWIP1. AtRanGAP1-GFP and RFP-Myc-AtSUN2 were co-expressed with either

AtWIP1 or AtWIP1XT in N. benthamiana leaves. Proteins were extracted from these two samples and immunoprecipitated with anti-GFP antibody. The co-IP of AtWIP1 and

AtWIP1XT was detected by the anti-WIP1 antibody raised against the N-terminus of

AtWIP1 (Xu et al., 2007). As shown in Figure 2.4E, both AtWIP1 and AtWIP1XT could be precipitated by AtRanGAP1-GFP. However, the co-immunoprecipitated RFP-Myc-

AtSUN2, detected by anti-Myc antibody, was only present in the AtWIP1 co-expressed sample, indicating that AtSUN2 indirectly interacts with AtRanGAP1 through AtWIP1.

Therefore, the SUN-WIP interaction functions in anchoring RanGAP to the NE in plants.

In mammals, during mitosis, the RanGAP anchor RanBP2 is re-located to the

42 kinetochores (KT) and is required for the KT localization of RanGAP (Joseph et al.,

2004). In plants, no RanBP2 homologs have been found. Like mammalian RanGAP,

AtRanGAP1 is also localized to the KT. In addition, it has plant-specific mitotic localizations—the preprophase band (PPB) which proceeds to the cortical division site

(CDS) during cell division. However, these mitotic RanGAP localizations are unchanged in the wip1-1 wip2-1 wip3-1 mutant (Xu et al., 2007), suggesting that WIP proteins are not the mitotic anchors of plant RanGAP. To test if plant SUN proteins could potentially mitotically anchor RanGAP through interactions with other unidentified plant KASH proteins, we examined the localization of AtSUN1 and AtSUN2 YFP fusions from prophase to metaphase in tobacco BY-2 suspension culture cells (Figure 2.7C). AtSUN1 and AtSUN2 were absent from PPB, CDS, and KTs, suggesting that they are unlikely involved in the mitotic localization of plant RanGAP.

2.4.6 AtSUNs and AtWIPs are required for maintaining an elongated nuclear shape

in epidermal cells wip1-1 wip2-1 wip3-1 or sun1-KO sun2-KD plants show no obvious developmental defects under laboratory conditions. The only phenotype observed in a previously reported AtSUN1/AtSUN2 double mutant is the increase in circularity of root hair cell nuclei (Oda and Fukuda, 2011). Thus, we investigated nuclear morphology in both sun1-

KO sun2-KD and wip1-1 wip2-1 wip3-1 plants. Since trichome nuclei and a portion of leaf epidermal cell nuclei are also elongated, we included these two cell types in this examination.

The fully expanded new leaves from plants before bolting were DAPI stained and observed under a fluorescence microscope. Our results show that the nuclei of leaf epidermal cells and trichomes are significantly less elongated in both mutants (Figure

2.5A). To compare the nuclear shape quantitatively, a ratio of nuclear width and length is used as an index to reflect the nuclear circularity. More elongated nuclei will have a lower circularity index, while rounder nuclei will have a circularity index closer to 1. For consistency, the maximum cross section or a z-stacked image of each nucleus was used to calculate the circularity index. As shown in Figure 2.5B (top two histograms), wild type has a significantly lower circularity index than both mutants, and the difference was confirmed by t-test (P<0.01, for leaf epidermal cells n=60, for trichomes n=20).

To observe root hair nuclei, roots from 7-8 day old seedlings were DAPI stained. In wild- type root hairs, DAPI staining revealed a super-elongated nuclear shape: a “pod” shape with two stretched thin tails extending from the poles (Figure 2.5A, third panel). This super-elongated nuclear shape has not yet been reported, but might be a variant of the fragmented and bi-lobed root hair nuclei observed by Chytilova et al. (2000). We then observed the root hair nuclear shape in the sun1-KO sun2-KD and wip1-1 wip2-1 wip3-1 mutants. As shown in Figure 2.5A, not only were the “pods” less elongated, but the extending tails were also lost. To quantitatively compare this nuclear shape change, we used the maximum length of a nucleus (including the tails) as an index. Figure 2.5B bottom histogram shows that wild-type nuclei are significantly more elongated than the nuclei in both mutants (t-test, P<0.01, n=55). To exclude that the process of DAPI

44 staining caused distortion of these nuclei or that DAPI would not reveal the full extent of the nucleoplasm, we also examined a GFP-NLS-GFP line (NLS fused with double GFP), using confocal microscopy. The bottom panel of Figure 2.5A shows that the same nuclear shape changes are observed when nuclei are visualized by the GFP marker that represents the entire nucleoplasm. In addition, Figure 2.5C shows a WPP-GFP line (WPP domain of

AtRanGAP1 fused with GFP, serving here as an NE marker), illustrating that the super- elongated nuclear shape was also observed when the NE was labeled. Together, these data suggest that AtWIPs and AtSUNs are required to maintain the elongated or super- elongated nuclear shape in three different types of plant epidermal cells.

In animals, a predominant function of SUN-KASH bridges is in the regulation of nuclear positioning and nuclear migration, through interactions with the cytoskeleton (Crisp et al., 2006). To investigate if the AtSUN-AtWIP interactions described here also affect nuclear positioning in Arabidopsis, two characterized nuclear positioning events were assayed. First, the position of the root hair nucleus is held at a specific distance from the growing tip by a process involving actin (Ketelaar et al., 2002). And second, the nucleus in a leaf hair (trichome) migrates to a fixed position close to the first branch point of the trichome cell (Folkers et al., 1997). Both processes were unchanged in both sun1-KO sun2-KD and wip1-1 wip2-1 wip3-1 (Figure 2.8). This indicates that AtSUN-AtWIP interactions do not contribute significantly to nuclear mobility and positioning in root and leaf hairs.

2.5 Discussion

2.5.1 Interactions between AtSUN and AtWIP

In this study, a combination of co-immunoprecipitation and fluorescence recovery after photobleaching experiments demonstrated that AtWIP1, AtWIP2, and AtWIP3 interact with AtSUN1 and AtSUN2 at the plant NE (Figure 2.2 and Figure 2.3). The PNS tail of

AtWIP1, especially the “VPT” motif, is required for the interaction, similar to the interactions between mammalian Nesprins and SUNs (Crisp et al., 2006; Padmakumar et al., 2005; Stewart-Hutchinson et al., 2008). Surprisingly, the PNS tail of WIP1 is only 9 amino acids long. The PNS tail of Homo sapiens Nesprin-2 is 30 amino acids long, and its C-terminal 14 amino acids are the shortest fragment able to bind the SUN domain

(Sosa et al., 2012). The ability of Arabidopsis SUN proteins to bind a very short PNS tail might be connected to the presence of a stretch of additional conserved residues in plant

SUN domains (Figure 2.6), however further work is required to understand the nature of the unusual plant SUN-dependent NE bridging complexes.

2.5.2 Function of the AtSUN-AtWIP complex in RanGAP NE anchoring

The NE localization of AtWIP1 is reduced in a sun1-KO sun2-KD mutant (Figure 2.4 A and B), suggesting that the localization of AtWIPs depends on AtSUNs, analogous to the animal KASH proteins. Consistent with these findings, the NE localization of

AtRanGAP1 is also reduced in undifferentiated root cells of the sun1-KO sun2-KD mutant (Figure 2.4 C and D), indicating that plant SUN proteins play a role in RanGAP-

NE association by forming a SUN-WIP-RanGAP complex. The existence of this complex is supported by co-immunoprecipitation data (Figure 2.4E). AtRanGAP1 is currently the only confirmed NE bridge cytoplasmic partner that is not apparently associated with elements of the cytoskeleton. RanGAP is the GTPase activating protein for the small

GTPase Ran. RanGTP hydrolysis is an important step in nucleocytoplasmic trafficking and implied in the release of cargo from export receptors after exiting from the nuclear pore at the cytoplasmic surface (Guttler and Gorlich, 2011). Both metazoan and plant

RanGAP is associated with the outer NE, but the mechanisms differ. The SUN-WIP-

RanGAP complex appears to be specific for plants. Mammalian nuclear pore-associated

RanGAP is in a complex containing the SUMO E3-ligase nucleoporin RanBP2,

SUMOylated RanGAP and the E2 SUMO-conjugating enzyme UBC9 (Reverter and

Lima, 2005). Plant RanGAP has no similarity to the SUMOylated domain and no

RanBP2 homolog exists in plants. In yeast and filamentous fungi, RanGAP is a cytoplasmic protein with no specific association with the NE (Hopper et al., 1990; Rose and Meier, 2001).

The finding that plants have recruited SUN-dependent NE bridges to anchor RanGAP to the NE suggests that functions for SUN-dependent NE bridges complexes beyond linking the nucleus and the cytoskeleton should be expected. Thus, novel, cytoskeleton-unrelated binding partners might also exist for opisthokont KASH proteins. One such KASH protein candidate might be C. elegans KDP-1, which functions in cell cycle progression

47 from late S to M phase, and for which the cytoplasmic partner is currently unknown

(McGee et al., 2009).

2.5.3 Function of the AtSUN-AtWIP complex in nuclear shape determination

Here we showed that the elongated epidermal nuclear shape is regulated by the AtSUN-

AtWIP complexes (Figure 2.5). A recent study suggests that the elongated nuclear shape is regulated by myosin XI-I which may interact with the AtSUN-AtWIP complex through the WPP domain-interacting tail-anchored protein 1 (AtWIT1) and AtWIT2. The details will be discussed in CHAPTER 3.

Nonetheless, the function of the elongated nuclear shape in plant epidermal cells is currently unknown, but it is conceivable that it might accommodate protection against environmental mechanical stress or endogenous shearing forces of cytoplasmic streaming. Indeed, mammalian linkers of nucleoskeleton to cytoskeleton complex (LINC) components are implied in reduced nuclear mechanotransduction and the activation of mechanosensitive genes (Lammerding et al., 2006; Lammerding et al., 2005;

Lammerding et al., 2004). In addition, nuclear shape changes have been correlated with both human disease and cell aging (Dahl et al., 2008; Webster et al., 2009).

Laminopathies, such as Hutchinson-Gilford progeria syndrome are diseases caused by mutations in type A Lamins. The nuclei of laminopathy patients often bear lobulated and invaginated nuclear shapes and changes in chromatin organization, leading to alteration in nuclear rigidity and sensitivity to mechanical stress. Similar nuclear shape changes occur during aging in C. elegans (Haithcock et al., 2005) and human cells (Scaffidi and

Misteli, 2006). In plants, nuclear shape changes have also been observed in the leaf epidermal cells of Arabidopsis nucleoporin nup136 mutants (Tamura et al., 2010) and the root cells of Arabidopsis bearing mutations in long coiled-coil NE protein LITTLE

NUCLEI1, 2, and 4 (Dittmer et al., 2007; Sakamoto and Takagi, 2013). It will be important to determine the interaction of these proteins with the SUN-WIP complex described here and to develop assays that test the effect of mechanical stress on nuclear function in plant epidermal cells.

In summary, we propose that WIP proteins are plant-specific KASH-like proteins that interact with plant SUN proteins at the NE in a mechanism similar to the known opisthokont SUN-KASH interactions. WIPs are anchored to the ONM by INM SUNs through interactions between the PNS tail of WIPs and the SUN domain of SUNs. This complex may be enriched close to the nuclear pores, where AtWIPs are concentrated (Xu et al., 2007). RanGAP is tethered to the ONM by WIPs and is in a complex with SUNs.

The nucleoplasmic N-terminus of SUNs may bind other INM proteins, and WIPs may recruit other cytoplasmic proteins involved in maintaining the elongated epidermal nuclear shape. Prior to this study, all known cytoplasmic connections of SUN-dependent

NE bridges were to the cytoskeleton, with functions in nuclear positioning or chromosome movement. The data presented here indicate that structurally and functionally diverged NE bridging complexes can be found in non-opisthokont branches of the tree of life.

Figure 2.1 Structural and sequence similarity between KASH domains and the PNS tail of AtWIP1. C-termini of animal and fungal KASH proteins are aligned with the C-terminus of AtWIP1. Extent of the TMD and the PNS tail are indicated below the alignment. Two-letter abbreviations of species names are as indicated in the legend of Figure 2.6. ClustalX color is assigned to the alignment for convenient comparison.

Figure 2.2 Characterization of AtSUN-AtWIP interactions. (A) Domain organization of AtWIP1 and mutant derivatives. AtWIP1 has an N-terminal domain with unknown function (cyan), an NLS (blue), a CCD binding AtRanGAP1 (red), a predicted TMD (yellow), and a PNS tail (shown in residues). (B) Domain organization of AtSUN2 and deletion constructs. AtSUN2 has an N-terminal domain with unknown function (cyan), a NLS (blue), a TMD (yellow), a unknown domain (white), a CCD (red), and a SUN domain (here split to an N-terminal part (green) and a C-terminal part (orange)) Figures in (A) and (B) are drawn to scale. (C) AtWIP1 interacts with AtSUN2 through its PNS tail. (D) AtWIP1, AtWIP2, and AtWIP3 interact with AtSUN2. (E) AtWIP1 interacts with AtSUN1 through its PNS tail. (F) AtWIP1, AtWIP2, and AtWIP3 interact with AtSUN1. (G) AtSUN2 interacts with AtWIP1 through its SUN domain. In (C) - (G), GFP- or CFP-tagged proteins were immunoprecipitated (IP) and detected by anti-GFP antibody. RFP-Myc- or Myc-Flag-tagged proteins were detected by anti-Myc antibody and RFP-Flag tagged proteins were detected by anti-Flag antibody. The input/IP ratio is 1/10. Numbers on left indicate mol wt in kD.

Figure 2.3 FRAP analysis of the interaction between AtWIP1 and AtSUN1. (A) Recovery curves of GFP-AtWIP1, GFP-AtWIP1VVPT, and GFP-TDFAtWIP1. (B) Recovery curves of GFP-AtWIP1 co-expressed with RFP-Flag-AtSUN1 or RFP-Flag-AtSUN1∆NSUN. (C) Recovery curves of GFP-AtWIP1 co-expressed with RFP-Myc-AtSUN2 or RFP- Myc-AtSUN2∆NSUN. (D) Recovery curves of GFP-AtWIP1VVPT co-expressed with RFP-Flag-AtSUN1 or RFP-Myc-AtSUN2. (E) Recovery curves of GFP-TDFAtWIP1 co- expressed with RFP-Flag-AtSUN1 or RFP-Myc-AtSUN2. In (A) – (E), error bars represent standard errors (for GFP-AtWIP1 in (C), n=60, for all others, n=30). Asterisks at the end of each curve indicates significant statistical difference of the maximum recovery compared to the green curve in each figure (t-test, P<0.01). Otherwise, no statistical difference has been observed (t-test, P>0.05) 52

Figure 2.4 AtSUNs are required for targeting AtWIP1 and AtRanGAP1 to the NE. (A) GFP- AtWIP1 or GFP-AtWIP1VVPT signal in undifferentiated root cells (top panel) and corresponding intensity profiles along the magenta arrows (bottom panel). C1, C2, cytoplasmic intensity 1, cytoplasmic intensity 2, respectively; N1, N2, nuclear intensity 1, nuclear intensity 2, respectively. Scale bars equal 5m. (B) NLI ([N1+N2]/[C1+C2]) calculated using the intensities measured as shown in (A). Asterisks indicate significant statistical difference between compared lines (t-test, P<0.01, n=50). Error bars represent standard errors. (C) AtRanGAP1-GFP signal in undifferentiated root cells (top panel) and corresponding intensity profiles along the magenta arrows (bottom panel). Scale bars equal 5m (D) NLI calculated as described in (B), using intensities measured as in (C). Asterisks indicate significant statistical difference between compared lines (t-test, P<0.01, n=55). Error bars represent standard errors. (E) AtSUN2, AtWIP1, and AtRanGAP1 are in the same complex. AtRanGAP1-GFP was immunoprecipitated and detected by anti-GFP antibody. AtWIP1 and RFP-Myc-AtSUN2 were detected with anti- AtWIP1 antibody and anti-Myc antibody, respectively. Numbers on the left indicate mol wt in kD.

Figure 2.5 Nuclear shape change in epidermal cells of sun1-KO sun2-KD and wip1-1 wip2-1 wip3-1 plants. (A) Comparison of nuclear shapes in trichomes, leaf epidermal cells, and mature root hair cells of wild type, sun1-KO sun2-KD, and wip1-1 wip2-1 wip3-1. Nuclei were DAPI stained, and mature root hair nuclei images in the last row are confocal maximum intensity projections using GFP-NLS-GFP as a nuclear marker. Scale bars equal 10 µm. (B) Quantitative comparison of nuclear shape changes shown in (A). Asterisks indicate significant statistical difference (t-test, P<0.01), compared to wild type. Error bars represent standard errors. (C) Confocal maximum intensity projection showing a super-elongated nucleus in a wild-type mature root hair using WPP-GFP as a NE marker. Scale bars equal 10μm.

Figure 2.6 Plant-specific conserved residues in plant SUN domains revealed by the alignment of SUN domains from different species. To identify plant SUN proteins, the protein sequence of AtSUN1 (At5g04990) was used for a BLAST search (http://blast.ncbi.nlm.nih.gov) against the green-plant (taxid: 33090) non-redundant protein sequence database (NCBI). Protein sequences showing high similarities to the SUN-domain region and Expect value <0.0001 were selected. Sequences of SUN proteins immediately after the predicted coiled-coil domain were aligned using ClustalW2. (http://www.ebi.ac.uk/Tools/msa/clustalw2/). Plant SUN domains are identified by species common name followed by the NCBI GenInfo Identifier number: poplar, Populus trichocarpa; ricinus, Ricinus communis; grape, Vitis vinifera; sorghum, Sorghum bicolor; maize, Zea mays; rice, Oryza sativa; lycophyte, Selaginella moellendorffii; moss, Physcomitrella patens. Non-plant SUN proteins are identified by 2- letter abbreviations of species names followed by the NCBI GenInfo Identifier number: Sp, Schizosaccharomyces pombe; Sc, Saccharomyces cerevisiae; Dd, Dictyostelium discoideum; Ce, Caenorhabditis elegans; Dm, Drosophila melanogaster; Mm, Mus musculus; Hs, Homo sapiens. The alignment was assigned with the ClustalX color scheme using Jalview. The plant SUN domains were group colored in Jalview (indicated by a light green frame), which highlights the plant-specific residues (denoted by red asterisks on top of the alignment). The proposed range of plant SUN domains is indicated by the top dark-green line, and the range of non-plant SUN domains is indicated by the bottom red line. The dark green arrow head denotes the break point of the Arabidopsis NSUN and CSUN domains used in this study.

Figure 2.6 Plant-specific conserved residues in plant SUN domains revealed by the alignment of SUN domains from different species.

Figure 2.7 AtWIP1 and AtSUN localization and characterization of sun1-KO sun2- KD. (A) RFP-Flag-AtSUN1 was co-expressed in N. benthamiana leaves with GFP- AtWIP1 (first row), GFP-AtWIP1∆VVPT (second row), and GFP-TDFAtWIP1 (third row). RFP-Flag-AtSUN1∆NSUN was co-expressed with GFP-AtWIP1 (fourth row). These samples were used for FRAP assay (as illustrated in the bottom row, for results see Figure 2.3). The white circle indicates the region that was photobleached and monitored for recovery. Scale bars equal 10 µm. (B) Semi-quantitative RT-PCR showing the mRNA levels of AtSUNs in sun1-KO sun2-KD. Total RNA was extracted from mature (before bolting) plants and used as templates for generating cDNA with M-MuLV reverse transcriptase and random hexamer primers (NEB Protoscript M-MuLV Taq RT-PCR kit). PCR products were obtained using primers amplifying AtSUN1 CDS (Lane A, forward primer: 5’-ATGTCGGCATCAACGGTGTCG-3’; reverse primer: 5’- TTATTCACTTTCAGGTGAAGAGTCCTG-3’), AtSUN2 CDS (Lane B, forward primer: 5’-ATGTCGGCGTCAACGGTGTC-3’; reverse primer: 5’- TCAAGCATGAGCAACAGAGAC-3’), and PP2A (Lane C, forward primer: 5’- ATGTCTATGGTTGATGAGCC-3’; reverse primer: 5’- GCTAGACATCATCACATTGTC-3’). The results show that PP2A (lane C) mRNA is present in both wild-type and sun1-KO sun2-KD samples and has similar levels in both samples. AtSUN1 (lane A) mRNA is present in the wild-type sample but not in the sun1- KO sun2-KD sample. AtSUN2 (lane B) mRNA is present is both wild-type and sun1-KO sun2-KD samples but with a lower level in the sun1-KO sun2-KD samples, indicating that sun1-KO sun2-KD is an AtSUN2 knock-down line. PCR without templates (last three lanes) or PCR using total RNA as templates (Lane D) resulted in no PCR products. (C) AtSUN1 and AtSUN2 mitotic localization in tobacco BY-2 cells. BY-2 cells were transformed and synchronized according to former describe protocols (Evans and Graumann, 2011). AtSUN1-YFP or AtSUN2-YFP (shown in green) were stably expressed in BY-2 cells and chromatin labeled with Draq5 (Biostatus, shown in magenta). Different mitotic stages are shown and AtSUNs were absent from the preprophase band (prophase), cortical division zone (prometaphase/metaphase), or kinetochores (prometaphase/metaphase), three sites of RanGAP localization in mitotic plant cells (Xu et al., 2007; Xu et al., 2008). Scale bars equal 10 µm.

Figure 2.7 AtWIP1 and AtSUN localization and characterization of sun1-KO sun2-KD.

Figure 2.8 Nuclear position in root hairs and trichomes is not affected in sun1-KO sun2-KD, and wip1-1 wip2-1 wip3-1. (A) Trichome nuclear position of wild type, sun1- KO sun2-KD, and wip1-1 wip2-1 wip3-1 were observed from fully expanded young leaves of 25-day old plants. The nuclei are highlighted by red dotted circles. Scale bars equal 20 µm. In wild type, the trichome nuclei are close to the first branch point (Folkers et al., 1997), and this position is not affected in sun1-KO sun2-KD and wip1-1 wip2-1 wip3-1 as shown by the relative nuclear position histogram (calculated by the distance from the nucleus to the baseline of a trichome divided by the distance from the second branch point to the baseline). Student t-test show no significant differences among wild type, sun1-KO sun2-KD, and wip1-1 wip2-1 wip3-1 (P>0.05, n=50). (B) Nuclear position of root hairs was observed using 11-day old seedlings. In wild type, the nucleus immigrates to the growing root hair and keeps a fix distance from the tip (Chytilova et al., 2000), which is unchanged in sun1-KO sun2-KD and wip1-1 wip2-1 wip3-1. Scales bars equal 20 µm.

Figure 2.8 Nuclear position in root hairs and trichomes is not affected in sun1-KO sun2-KD, and wip1-1 wip2-1 wip3-1.

CHAPTER 3

Efficient plant sperm cell delivery requires vegetative nuclear movement mediated

by WPP domain-interacting tail-anchored proteins

3.1 Abstract

In flowering plants, the sperm cells (SCs) are delivered to ovules by pollen tubes which extend over a great distance. The movement of the SCs is tightly associated with the vegetative nucleus (VN), which is proposed to be essential for efficient SC delivery.

However, no evidence has been shown to support this hypothesis, and little is known about the molecular mechanism regulating the VN movement. Here, we identified the

Arabidopsis thaliana WPP domain-interacting tail-anchored protein 1 (WIT1) and WIT2 and their binding partners WPP domain-interacting protein 1 (WIP1), WIP2, and WIP3, as key players in VN movement. Loss of either WIT1 and 2 or WIP1, 2 and 3 impaired the VN movement and resulted in inefficient SC-to-ovule delivery. We further showed that the VN movement was not regulated by the recently identified WIT-myosin XI-I complexes.

3.2 Introduction

Unlike animals whose sperm cells (SCs) travel towards egg cells through self-powered flagellum propelling, flowering plants utilize pollen tubes to deliver SCs to ovules. Pollen is generated by pollen mother cells which undergo meiosis to generate microspores. A microspore continues to divide asymmetrically to produce mature bicellular pollen that contains a generative cell (GC). In another type of pollen formation, the GC undergoes one more round of mitosis to produce two SCs, forming a mature tricellular pollen grain.

The vegetative nucleus (VN) of pollen and the GC/SC are usually closely associated with each other in both pollen grains and pollen tubes (for recent review see McCue et al.,

2011). In many plant species, a connection between the VN and the GC/SC has been identified, and therefore, they are termed as “male germ unit” (MGU) (Dumas et al.,

1985; McCue et al., 2011). The MGU was proposed to be important for successful fertilization by signal transduction between the VN and SC (Dumas et al., 1985) or by efficient and simultaneous SC delivery (Russell and Cass, 1981).

In growing pollen tubes of many angiosperm species, the VN precedes (closer to the growing pollen tube tip) the GC/SC (Heslop-Harrison and Heslop-Harrison, 1989b;

Lalanne and Twell, 2002; McCue et al., 2011). However, the order can be more complicated in some species. During Amaryllis vittata Ait. pollen tube growth, either the

VN or the GC enters the tube first, but afterwards, the VN always precedes the GC.

When the GC undergoes mitosis, the position of VN and GC starts shifting. A similar process was also observed during Nicotiana tabacum pollen tube growth, where the VN

63 was ahead of the GC in the beginning and later became proximal to the dividing GC

(Palevitz, 1993). The function of this VN reposition during GC division is unknown.

The molecular mechanism underlying the MGU movement has been studied for decades, but it still remains an enigma. Depolymerizing microtubules in bicellular pollen tubes using oryzalin or colchicine resulted in a spectrum of defects depending on the concentration: it restricted VN and GC movement (Astrom et al., 1995; Kaul et al.,

1987), impaired the order of the VN and GC in pollen tubes (Heslop-Harrison et al.,

1988), separated the VN further from the GC (Heslop-Harrison et al., 1988), increased the distance between the leading nucleus and the growing tube tip (Heslop-Harrison et al., 1988), affected microfilament organizations (Kaul et al., 1987), and delayed pollen tube growth. These results suggest that the microtubule network is responsible for MGU movement. On the other hand, anti-myosin antibodies labeled the VN periphery of several plant species (Heslop-Harrison and Heslop-Harrison, 1989c; Miller et al., 1995;

Tirlapur et al., 1995). Depolymerizing microfilaments also led to abnormal VN and GC positioning in pollen tubes, suggesting that the microfilament network may also be involved in MGU movement. In spite of these studies, no solid evidence has been presented to assign any genes to the MGU movement. Our limited understanding of the

MGU movement is in great contrast to its importance for pollen tube growth and fertilization.

In animals, nuclear movement is usually regulated by the linker of the nucleoskeleton and cytoskeleton (LINC) complex formed by Klarsicht/ANC-1/Syne-1 homology (KASH)

64 proteins and Sad1/UNC-84 proteins. KASH proteins at the cytoplasmic side can recruit motor proteins and transfer forces to the nucleus through LINC complexes (Gundersen and Worman, 2013; Razafsky and Hodzic, 2009; Starr and Fridolfsson, 2010). In plants, the WPP domain-interacting proteins (WIPs) were recently identified as analogs of

KASH proteins. Arabidopsis thaliana WIP1, WIP2, and WIP3 form bridging complexes with SUN1 and SUN2 at the NE (Zhou et al., 2012a). These SUN-WIP bridges are important for anchoring Ran GTPase activating protein 1 (RanGAP1) to the NE and maintaining elongated nuclear shape of epidermal cells (Zhou et al., 2012a). WIP1,

WIP2, and WIP3 also interact with WPP domain-interacting tail-anchored protein 1

(WIT1) at the NE (Zhao et al., 2008). WIT1 and WIT2 are additional plant RanGAP NE anchors that act semi-redundantly with WIP1, 2, and 3(Zhao et al., 2008). A recent study reported that WIT1 and WIT2 recruit myosin XI-I to the NE and that WIT1/WIT2- myosin XI-I complexes regulate the elongate nuclear shape in roots and the nuclear movement in both roots and leaf mesophyll cells, suggesting a potential role of WIT1 and

WIT2 in nuclear movement (Tamura et al., 2013).

Here, we report that WIT1 and WIT2 regulate the movement of the VN in germinating pollen tubes synergistically with WIP1, WIP2, and WIP3. Defects of VN movement impair SC delivery and lead to considerable seed loss. This function of the WIT and WIP protein families is uncoupled from the nuclear shape determination and the regulation of nuclear movement involving myosin XI-I. We show that 1) unlike for VN movement, in

65 epidermal cells WIT2 but not WIT1 played a key role in nuclear shape determination and

2) both WIT2 and myosin XI-I mutants had no significant seed loss phenotype.

3.3 Material and Methods

3.3.1 Plant materials

Arabidopsis (Columbia ecotype) plants were grown at 25⁰C in soil under 16-h light and

8-h dark or on MS (Caisson laboratories) with 1% sucrose plates under constant light.

Wit1-1 wit2-1 was reported by Zhao et al. (2008) and wip1-1 wip2-1 wip3-1 was reported by Xu et al. (2007). Crosses between these two mutants were performed to obtain the wip1-1 wip2-1 wip3-1 wit1-1 wit2-1 (wifi) mutant. Kaku1-4 (SALK_082443C) was reported by Tamura et al. (2013) and the seeds were obtained from the Arabidopsis

Biological Resource Center. The sun1-KO sun2-KD mutant (SALK_123093 for sun1-KO and SALK_049398 for sun2-KD) was a gift from Dr. Susan Armstrong (University of

Birmingham, UK) and has been introduced in CHAPTER 2.

3.3.2 Arabidopsis cDNA synthesis

Arabidopsis whole seedlings (7 day old) grown on MS with 1% sucrose plates were harvested and ground to power in liquid nitrogen. One ml TRIzol (Life Technologies) was added to 500 µl of tissue powered, mixed well, and left at room temperature for 5 min. 200 µl chloroform was then added to the mixture, mixed well, and left at room temperature for 5 min. After centrifugation at 20000g, 4ºC for 15 min, the supernatant was used for RNA preparation using RNeasy Plant Mini Kit (Qiagen). The cDNA was synthesized from the extracted RNA using the SuperScript® III First-Strand Synthesis

System (Life Technologies).

3.3.3 Constructs

The hygromycin B resistance cassette was amplified by PCR from the vector pH2GW7

(Karimi et al., 2002) using 5’-AATGAATTCATCAGCTTGCATGCCGGTCGATC-3’ and 5’-GCTGAATTCATCATACATGAGAATTAAGGGAGTC-3’ (the EcoRI site is underlined), digested by EcoRI, and then ligated with the EcoRI-digested binary vector pPZP-RCS2 (Goderis et al., 2002). After confirmation by sequencing, the pPZP-RCS2-

Hyg vector was obtained. The GFP-gateway-35ST (35S terminator of Cauliflower

Mosaic Virus) cassette was amplified by PCR using 5’-

TATGGCGCGCCACGTGAGCAAGGGCGAGGAGCTGTTC-3’ (the AscI site is underlined) and 5’-CCGGGGATCCTCTAGAGGGCC-3’ (the XbaI site is underlined), digested by AscI and XbaI, and ligated with the AscI/XbaI-digested pPZP-RCS2-Hyg.

After confirmation by sequencing, the pHOAG vector was obtained.

The WIT1 promoter sequence (~ 2.1kb upstream of the start codon of WIT1) was amplified by PCR from Arabidopsis (ecotype Columbia) DNA using 5’-

ATCGAGCTCCAATGGGTCCTGTGTTGGTCCACG-3’ (SacI site is underlined) and

5’-CATCTTTCAATATAACTGCAACAGAGAAAGTA-3’, digested by SacI, and ligated with the SacI/PmlI-digested pHOAG. After sequencing, the pHWIT1proAG vector was obtained. WIT1 CDS cloned in pENTR/D-TOPO vector (Life Technologies) was described by Zhao et al. (2008), and was moved to pHWIT1proAG by an LR reaction (Life Technologies) to obtain the WIT1pro::GFP-WIT1 construct. The WIP1 promoter (~ 2.6kb upstream of the start codon of WIP1) was amplified by PCR from

Arabidopsis DNA using 5’-GGAAGGCGCGCCCACCGTTATGACTCG-3’ (AscI site is underlined) and 5’-CATTGACTCCACAAAAAAATCTATC-3’, digested by AscI, and ligated with the AscI/PmlI-digested pHOAG. After confirmation by sequencing, the pHWIP1proAG vector was obtained. WIP1 CDS cloned in pENTR/D-TOPO vector was described by Xu et al. (2007), and was moved to pHWIP1proAG by an LR reaction to obtain the WIP1pro::GFP-WIP1 construct. The WIT2 promoter (~2.2kb upstream the start codon of WIT2) was amplified by PCR from Arabidopsis DNA using 5’-

GGCCCGGCGCGCCACTGATGAATCATTCACCAAGAGTGGT-3’ and 5’-

CTCGCCCTTGCTCACCATTGACTCCACAAAAAAATCTATC-3’, and cloned into the PmlI-digested pHOAG by In-fusion (Clontech) recombination. After confirmation by sequencing, the pHWIT2proAG vector was obtained. WIT2 CDS was cloned from

Arabidopsis whole seedling cDNA (see 3.3.2) using 5’-

GGGGACAAGTTTGTACAAAAAAGCAGGCTTTATGGAGGAAATCATTAGGGA

GGAC-3’ and 5’-GGGGACCACTTTGTACAAGAAAGCTGGGTA

TTAATAAGTCACACCAAAGAATGAA-3’, cloned into pDONR221 (Life

Technologies) by a BP reaction (Life Technologies). After sequencing the PCR product and the sequence cloned into pDONR221, it was found that the T182-C562 fragment of the WIT2 CDS was constantly (3 independent cloning events, and same results) lost in E. coli, resulting in a sequence encoding gene model At1G68910.3 with G62-L188 deleted).

This truncated WIT2 was named as WIT2*. WIT2* CDS was then moved from pDONR221 to pHWIT2proAG by an LR reaction to obtain the WIT2pro::GFP-WIT2*

69 construct. WIT2* CDS in pDONR221 was also cloned into pH7WGF2 by an LR reaction to obtain 35S promoter-driven GFP-WIT2*. WIP1, WIP2, and WIP3 CDS cloned in pENTR/D-TOPO were described by Xu et al. (2007), and were moved to pGWB21

(Nakagawa et al., 2007) by LR reactions to obtain 35S promoter-driven Myc-WIP1, Myc-

WIP2, and Myc-WIP3, respectively.

3.3.4 Generation of transgenic plants

Binary constructs were transformed to Agrobacterium strain ABI by triparental mating

(Wise et al., 2006).The VN marker, λN22 (an RNA binding protein) fused with mCherry and an NLS driven by the ubiquitin 10 promoter (VN-RFP) (Schönberger et al., 2012), and the SN marker, male-gamete-specific histone H3 fused with GFP driven by the

MGH3 promoter (SN-GFP) (Ingouff et al., 2007), used in this study are gifts by Dr. Keith

R. Slotkin. The VN-RFP and SN-GFP marker Agrobacterium strains and the wild-type

Arabidopsis line transformed with both markers are gifts by Dr. Keith R. Slotkin. Other transgenic Arabidopsis lines were obtained by Agrobacterium-mediated floral dip

(Clough and Bent, 1998).

3.3.5 Co-immunoprecipitation experiments

Agrobacterium cultures containing plasmids for expressing the proteins of interest were co-infiltrated into N. benthamiana leaves as described previously (Sparkes et al., 2006).

Plants were grown 3 days after infiltration, and leaves were collected and ground in liquid nitrogen into powders. All immunoprecipitation steps were performed at 4⁰C. For

70 details please see 2.3.4. Briefly, protein extracts were prepared in NP-40 buffer containing 50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.5% NP-40, 1 mM EDTA, 3 mM

DTT, 1 mM PMSF, and 1% protease inhibitor cocktail (Sigma-Aldrich). Protein complexes were immunoprecipitated from extracts by an anti-GFP antibody (ab290,

Abcam Cambridge) bound to A-sepharose beads (GE Healthcare). The immunoprecipitates were then analyzed by western blot with an anti-GFP (1:2000,

632569, Clontech) or an anti-Myc (1:1000, M5546, Sigma-Aldrich) antibody.

3.3.6 Hoechst 33342 staining and nuclear shape measurement

For Hoechst 33342 staining, a 4% paraformaldehyde solution containing 4 µM Hoechst

33342 was used to stain pollen grains and 7- or 8-day old Arabidopsis seedlings grown on MS plates for at least 20 min. After rinsing in PBS buffer (137 mM NaCl, 2.7 mM

KCl, 10 mM Na2HPO4, 2 mM KH2PO4, pH7.4), seedlings were mounted in PBS buffered and viewed under Nikon C90i microscope. The UV-2E/C filter cube (Nikon) was used for the imaging. For trichomes, leaves of 25-35-day old plants grow in soil were mounted in water and the trichomes at leaf edge were examined. The optically dense nuclei were directly imaged using bright field microscopy. Images were taken by a Nikon DS-Qi1Mc digital camera. The length and width of the nuclei were measured using the Nikon NIS-

Elements software.

3.3.7 In vitro pollen germination and Alexander staining

Pollen grains from the stamens of fully opened flowers were germinated on pollen germination medium containing 18% sucrose, 0.01% boric acid, 1 mM CaCl2, 1 mM Ca

(NO3)2, 1 mM MgSO4, and 0.5% agar. Several wild-type stigmas were placed adjacent to pollen grains to stimulate pollen germination (Qin et al., 2011). Alexander pollen staining was performed as described previously (Alexander, 1969).

3.3.8 Pollen tube confocal microscopy, length measurement, and kymograph

generation

A Nikon Eclipse C90i confocal microscope with minimum or medium aperture was used to collect images. For time course imaging of pollen tube growth, a z-stack was taken

(step size, 5 µm) every minute for 180 min. To maximize imaging speed, sequential imaging of GFP and RFP that blocks bleach through was not used, since the VN and SN can be easily distinguished by their size and shape.

Each channel of the pollen tube growth movie was first split in ImageJ. For every time point, the z-stack was converted to one frame using maximum projection. The GFP, RFP, and the transmitted light channels were then combined to one RGB color movie. If position shifting occurs, the StackReg plugin with the “translation” algorithm was used to correct the shift. Length was then measured using ImageJ. Kymographs were generated in ImageJ by first drawing a selection segmented line over the pollen-tube growth track, and then reslicing the stacks.

3.4 Results

3.4.1 WIT and WIP family proteins are required for full seed set

Arabidopsis encodes two WIT proteins—WIT1 and WIT2 (Zhao et al., 2008). Careful observation revealed that siliques of the wit1-1 wit2-1 double null mutant contain only about 50% seeds per silique when compared with wild type (Figure 3.1A). Although, the number of seeds per silique of wit1-1 and wit2-1 single mutants is statistically significantly different from wild type, the numbers are only slightly reduced when compared with wild type (Figure 3.1A), indicating that WIT1 and WIT2 are redundant for seed set. Seed loss was also observed in a wip1-1 wip2-1 wip3-1 triple knockout mutant but it is less severe (Figure 3.1A). The quintuple knockout mutant wip1-1 wip2-1 wip2-1 wit1-1 wit2-1 (wifi) has the most severe phenotype, indicating that the WIT and

WIP families synergistically affect seed production. WIT1 promoter-driven GFP-WIT1

(WIT1pro::GFP-WIT1) was transformed into wit1-1 wit2-1 and wifi. As shown in Figure

3.1A, this construct rescued two wit1-1 wit2-1 transgenic lines to the wit2-1 level (line 1 and line 3), partially rescued one wit1-1 wit2-1 line (line 2), and rescued two wifi transgenic lines to the wip1-1 wip2-1 wip3-1 level, confirming that this phenotype is linked to WIT1.

3.4.2 Loss of WITs and WIPs leads to defects of pollen fertility

As shown in Figure 3.1B, wit1-1 wit2-1 ovaries contain on average four ovules less than wild type (P<0.01, n=10, 1- tailed Student’s t-test), which is unlikely to cause the

73 significantly reduced seed production of wit1-1 wit2-1. We then performed reciprocal crosses between wit1-1 wit2-1 and wild type. As shown in Figure 3.1C, when pollinated with wild-type pollen, both wit1-1 wit2-1 and wild type have similar numbers of seeds per silique (P>0.05, 2-tailed Student’s t-test). In contrast, when pollinated with wit1-1 wit2-1 pollen, both wit1-1 wit2-1 and wild type have comparably reduced numbers of seeds per silique (P<0.01, 2-tailed Student’s t-test). Since wifi has the most severe phenotype, reciprocal crosses were also performed between wifi and wild type, and similar results were obtained (Figure 3.1C). These data indicate that WIT1 and WIT2, as well as at least one member of the WIP family, are involved in pollen fertility.

3.4.3 WIT1 and WIP1 are located at the vegetative NE in pollen grains

We then examined the localization of WIT1 and WIP1 in wild-type pollen grains using

GFP-tagged WIT1 and WIP1 driven by their native promoters. In wild-type pollen, both

GFP-WIP1 and GFP-WIT1 signals were strongly associated with the vegetative NE. In addition, GFP-WIT1 weakly labeled the sperm cell NE in some pollen grains (Figure 3.2

A and C). After five hours of in vitro growth of pollen tubes, GFP-WIP1 was still only visible at the vegetative NE, while WIT1 was now also weakly visible at the SC NE, in addition to the strong signal at the vegetative NE (Figure 3.2).

It has been reported that the WIT1 protein level is significantly reduced in the wip1-1 wip2-1 wip3-1 mutant, likely because the protein is destabilized/degraded in the absence of the WIP-WIT complex (Zhao et al., 2008). Therefore, we examined the GFP-WIT1 signal level in WIT1pro::GFP-WIT1 transformed wip1-1 wip2-1 wip3-1. The GFP-WIT1

74 signal is barely detectable in wip1-1 wip2-1 wip3-1 pollen (three lines were examined and a representative image is shown in Figure 3.2C), which coincides with the previous report. However, when WIT1pro::GFP-WIT1 was transformed into sun1-KO sun2-KD,

GFP-WIT1 localization in its pollen grains was indistinguishable from wild type (three lines were examined and a representative image is shown in Figure 3.2C). This is consistent with our observation that sun1-KO sun2-KD has negligible seed set defects

(Figure 3.1A). This also suggests that the reduced fertility phenotype of wip1-1 wip2-1 wip3-1 is to a large extent due to the loss of WIT proteins. Thus, we focused the subsequent experiments on WIT1 and WIT2.

Together, these data support a model of WIT1 and WIT2 association with the VN through one or more WIP family members and this association being required for full male fertility. The male fertility defect reported here is clearly different from the female gametophyte lethality caused by mutating RanGAP1 and RanGAP2 (Rodrigo-Peiris et al.,

2011).This suggests that maintaining pollen fertility is a function of WIT and WIP proteins independent of their reported role as RanGAP NE anchors.

3.4.4 WIT and WIP family proteins are required for proper nuclear movement in

pollen tubes

The morphology of wit1-1 wit2-1, wip1-1 wip2-1 wip3-1, and wifi anthers and pollen grains is normal and no significant amount of dead pollen was observed using Alexander staining (Figure 3.3A). The three nuclei—one VN and two sperm cell nuclei (SN)—are also normal in wit1-1 wit2-1, wip1-1 wip2-1 wip3-1, and wifi pollen grains (Figure 3.3B).

We then examined the nuclear movement of wit1-1 wit2-1 pollen grains germinated in vitro. A VN marker, mcherry fused with an NLS driven by the ubiquitin 10 promoter

(VN-RFP) (Schönberger et al., 2012), and a SN marker, male-gamete-specific histone

H3-GFP driven by the MGH3 promoter (SN-GFP) (Ingouff et al., 2007), were used.

As shown in Figure 3.4A, during wild-type pollen germination, the VN usually entered the pollen tube first (Figure 3.4D), when the pollen tube reached a certain length (we define the tube length when the first nucleus permanently enters the pollen tube as the

“entrance length”). The VN then kept a certain distance from the growing tip, while the two SN remained behind the VN and loosely followed the VN movement (Figure 3.4A).

Occasionally, when the VN arrived closely to the tip, a rapid backward movement was observed, which was followed by the SN, resulting in a serrate moving track (Figure

3.4A). Among 40 wild-type pollen grain germinations analyzed, the average entrance length was 35.5 µm (S.D.=7.8 µm, Figure 3.4C). We also followed the VN for 60 min after it entered the pollen tube and measured the VN-to-tip distance every minute. The

VN-to-tip distance oscillated as the pollen tubes grew, with its mean value being 27.1 µm

(S.D.=7.2 µm, 10 pollen tubes were measured, Figure 3.4E).

In contrast, when wit1-1 wit2-1 pollen tubes had reached their much greater entrance length (mean=95.9 µm, S.D.=29.5 µm, n=40, Figure 3.4C, Student’s t-test to wild type,

P<0.01), the two SN entered the pollen tube first in most cases, followed by the VN. This behavior was significantly different from wild type (Figure 3.4B and D, 2-tailed Fisher’s exact test, P<0.01). The two SN “led” the VN in the growing pollen tubes, even in

76 situations when the VN entered the pollen tubes first (Figure 3.4B and F), significantly different from wild type (2-tailed Fisher’s exact test, P<0.01). Measuring the leading-SN- to-tip distance every minute for 60 min after the two SN had entered the pollen tube, a mean value of 56.2 µm (S.D.=31.2 µm, 10 pollen tubes were measured, Figure 3.4E) was obtained, much longer than that of wild type (P<0.01, n=600, 2-tailed Student’s t-test).

The leading-nucleus-to-tip distances are significantly different between wild type and wit1-1 wit2-1 at every time point during the 60 min tracking (Figure 3.5, 2-tailed

Student’s t-test, P<0.05, n=10).

We then examined the nuclear position in in vitro germinated pollen tubes at 5 h, a time point when in vivo pollen tubes would start entering the micropyle (Faure et al., 2002). In nearly all wit1-1 wit2-1 pollen tubes, the two SN were still ahead of the VN, significantly different from wild type (Figure 3.4F, 2-tailed Fisher’s exact test, P<0.01). After 8 h of in vitro germination, a time point when the double fertilization starts in vivo, the SN still preceded the VN in wit1-1 wit2-1 pollen tubes, which is the opposite of what was observed in wild type (Figure 3.6). This implies that the VN movement is impaired during the entire length of pollen tube growth. A similar phenotype was also observed in

Hoechst 33342-stained wip1-1 wip2-1 wip3-1 and wifi pollen tubes after 5 h in vitro germination (Figure 3.4F, P<0.01, 2-tailed Fisher’s exact test to wild type). Consistently, the two WIT1pro::GFP-WIT1 transformed wit1-1 wit2-1 lines, whose seed loss phenotype was rescued (line 1 and line 3, Figure 3.1A) showed normal nuclear

77 positioning after 5 h of in vitro germination (Figure 3.4F, P>0.05, 2-tailed Fisher’s exact test to wild type).

3.4.5 Loss of WIT proteins impaired the SC-to-ovule delivery

The seed loss phenotype of wit1-1 wit2-1 may be due to the impaired movement of the

VN. To test this hypothesis, ~48 h after pollination ovaries were dissected. Two types of ovules were found in wit1-1 wit2-1 siliques—small unfertilized ovules whose central cell and egg cell were clearly visible and large, fertilized ovules with visible, developing embryos (Figure 3.7A). The number of large, fertilized ovules per ovary of wit1-1 wit2-1 was slightly lower than the seeds per silique of wit1-1 wit2-1, suggesting that a small fraction of the unfertilized ovules we scored would still become fertilized (compare

Figure 3.7B and Figure 3.1A). A small fraction of unfertilized ovules that later become fertilized was also found in wild type (compare Figure 3.7B and Figure 3.1A). Confocal imaging of the small, unfertilized ovules showed that the two SN were arrested in the vicinity of the ovules (30%), at the entrance of the micropyle (46%), or inside the ovules

(24%) (Figure 3.7), reflecting a severe SC delivery defect. In none of the analyzed ovules was a VN visible. The VN might never reach the vicinity of ovules or it might be degraded inside the pollen tube. Together, these data suggest that the observed defects in

VN movement lead to severe impairment in SC-to-ovule delivery.

3.4.6 VN movement is not mediated through the proposed SUN-WIP-WIT-myosin

XI-I complex

Recently, a forward genetic screen isolated myosin XI-I as a root nuclear shape regulator

(Tamura et al., 2013). WIT1 and/or WIT2 were reported to recruit myosin XI-I to the NE, and wit1-1 wit2-1 and myosin xi-i mutants have lost root nuclear elongation and display restricted nuclear movement. To test if the same complex is also responsible for movement of the VN, we examined whether kaku1-4, a reported myosin xi-i knockout mutant, has a seed set defect (Tamura et al., 2013). As shown in Figure 3.1A, kaku1-4 exhibits normal seeds per silique, suggesting that myosin XI-I is not necessary for VN mobility during pollen tube growth.

Surprisingly, further investigation showed that WIT1 and WIT2 were not equally determining epidermal nuclear morphology. Since root hair and trichome nuclei are most obviously elongated, they were used in this investigation. Because the root hair nuclei are extremely elongated and have a very variable shape and positioning (sometimes folding back on themselves when viewed in 3D), the simple total length of the longest extension of the root hair nucleus was used as an index. For trichome nuclei, the ratio of the width and length of the nucleus was used as a circularity index (the closer to 1, the more circular). For each line, 80 root hair nuclei and 50 trichome nuclei were measured. As shown in Figure 3.8 B and C, the circularity indices of wit2-1 in both cell types are significantly different from those of wild type (P<0.01, 2-tailed Student’s t-test) and are similar to those of wit1-1 wit2-1 (P>0.05, 2-tailed Student’s t-test), while the circularity

79 indices of wit1-1 are barely distinguishable from wild type. This indicates that WIT2, but not WIT1, is essential for nuclear elongation in epidermal cells.

To confirm the nuclear shape phenotype of wit2-1 was caused by the loss of WIT2, a partial WIT2 was cloned—the T182-C562 fragment of the WIT2 coding sequence was constantly lost in E. coli during cloning (at least 3 independent cloning trails), resulting in a sequence encoding gene model At1G68910.3 with G62-L188 deleted (WIT2*). GFP-

WIT2* driven by WIT2 promoter (WIT2pro::GFP-WIT2*) was transformed to wit2-1 and wit1-1 wit2-1. Three wit2-1 transgenic lines and one wit1-1 wit2-1 line were analyzed. As shown in Figure 3.8, GFP-WIT2* was localized to the NE and was able to partially complement the nuclear shape phenotype (compared with wit2-1 and wit1-1 wit2-1,

P<0.01, n=80 for root nuclei and n=50 for trichome nuclei, 2-tailed Student’s t-test). This indicates that WIT2* is partially functional and that the loss of nuclear elongation in wit2-1 and wit1-1 wit2-1 is based on the loss of WIT2.

The ability of WIT2* to bind WIP1, WIP2, and WIP3 was then tested. GFP-WIT2* was co-expressed with Myc-WIP1, Myc-WIP2, and Myc-WIP3, respectively, in N. benthamiana leaves. GFP-NLS-GFP-NES was co-expressed with Myc-WIP3 to serve as a non-specific GFP-fusion protein control. As shown in Figure 3.9, an anti-GFP antibody immunoprecipitated GFP-WIT2*, and all Myc-tagged WIPs were co- immunoprecipitated. In the negative control, GFP-NLS-GFP-NES did not precipitate

Myc-WIP3. These data suggest that WIT2 is able to interact with the SUN-WIP complexes.

However, neither wit2-1 nor kaku1-4 has a seed set defect comparable to wit1-1 wit2-1.

In addition, the seed set of sun1-KO sun2-KD is not considerably affected (Figure 3.1A), likely because the residual SUN2 expression is sufficient for anchoring the WIP-WIT complexes.

3.5 Discussion

3.5.1 The importance of MGU

The MGU has been observed in many plant species, and it has been proposed to be important for efficient SC delivery (Dumas et al., 1985; McCue et al., 2011; Russell and

Cass, 1981). Through our observation of in vitro geminated Arabidopsis pollen, the VN and SC do form a linked unit and move together in pollen tubes (Figure 3.4A). Even in the wit1-1 wit2-1 mutant whose VN seemingly loses the driving force, the VN still move together with the SCs (Figure 3.4B), reflecting a functional linkage between the VN and

SCs. That the impaired VN movement of wit1-1 wit2-1 led to inefficient SC delivery

(Figure 3.7) supports the proposed function of the MGU.

3.5.2 Driving forces of the MGU

During wild-type pollen tube growth, the VN enters the tube first when the pollen tube reaches a certain length (Figure 3.4C). In some cases, we observed that the VN entered the pollen tube first, but then reversed direction and moved towards the pollen grain.

Later, the VN “led” the movement of SCs with occasionally backward movement when the MGU was extremely close to the tip of the growing tube, resulting a serrated moving

81 curve in the kymograph (Figure 3.4A). Although the mobility of the VN is impaired in wit1-1 wit2-1 and the entrance length is longer than that of wild type, the distance- keeping ability of the MGU is not totally lost: the SCs “led” the movement of the MGU, and made back-and-forth adjustment according to the distance to the growing tube apex

(Figure 3.4B).

These observations indicate that 1) there is a main driving force transferred via WITs that moves the VN; 2) a secondary driving force facilitating the movement of the SCs should also exist, and 3) a signaling mechanism is present between the VN/SC and the growing tip regulating the two driving forces to maintain the nucleus-tip distance. As the VN and the two SN are connected, these two forces coordinate with each other to move the MGU forward. This “double-driving-force” hypothesis is consistent with a reported observation that the SC projection which links the VN exhibited somewhat independent movement to the overall momentum of SCs (Ge et al., 2011). In addition, the bidirectional movement of the MGU was proposed to be a result of the dynamic balance between tacropetal and basipetal forces (Heslop-Harrison and Heslop-Harrison, 1989a). Although the nuclear order defects and the increased leading-nucleus-to-tip distance of wit1-1 wit2-1 mimics the nuclear movement in the Galanthus nivalis L. pollen tubes after microtubule depolymerization (Heslop-Harrison et al., 1988), the question whether the MGU movement is regulated by the F-actin network or the microtubule network still needs further investigation.

3.5.3 Difference between nuclear movement in vegetative tissues and pollen tubes

Myosin XI-I was isolated as nuclear shape regulator in root cells and shown to regulate nuclear movement in root cells and leaf mesophyll cells (Tamura et al., 2013). This was reported to be achieved by binding WIT1 and WIT2, which may further link the SUN-

WIP NE bridges. Here, we show that WIT2, but not WIT1, is required for elongated nuclear shape in the sporophyte (Figure 3.8), while WIT1 and WIT2 act redundantly in controlling the VN movement in the male gametophyte. This suggests a different mechanism for the two processes. Consistently, kaku1-4 has no seed set defect (Figure

3.1A), indicating that the VN movement is not regulated by myosin XI-I. The most likely scenario is that WIT1 and WIT2 redundantly recruit another factor (likely another motor) to transport the VN in the male gametophyte.

Since both wit1-1 wit2-1 and wip1-1 wip2-1 wip3-1 have similar phenotypes and all

WITs interact with all WIPs, we propose that in pollen tubes WIT1 and WIT2 serve as an adaptor for the SUN-WIP NE bridges to recruit unknown factors to drive the movement of the VN.

Figure 3.1 Reduction in seed production based on loss of WITs and WIPs. (A) Number of seeds per silique compared between different mutants and lines. Asterisks represent statistical significance between the indicate data sets and the wild-type data set, while “o” represents no statistical significance was found (“*” represents 0.010.05, n=40, 2-tailed Student’s t-test). (B) Number of ovules per ovary of wild type and wit1-1 wit2-1. “**” represents statistical significance (P<0.01, n=10, 2-tailed Student’s t-test) (C) Number of seeds per silique after reciprocal crosses between wild type and wit1-1 wit2-1 or between wild type and wifi. “**” represents statistical significance (P<0.01) between compared data sets, while “o” presents no statistical significance was found (P>0.05, 2-tailed Student’s t-test, n for each data set was indicated in the corresponding column,). Error bars in all histogram represent S.D.

Figure 3.2 WIT1 and WIP1 subcellular localization in pollen grains and pollen tubes. (A) The localization of WIT1 and WIP1 in pollen grains was examined in WIT1pro::GFP-WIT1 transformed wild-type plants and WIP1pro::GFP-WIP1 transformed wild-type plants, respectively. GFP signal was imaged by confocal microscopy and Hoechst 33342 signal was imaged by fluorescence microscopy. WIT1 and WIP1 were both localized at the VN NE. In addition, WIT1 also weakly labeled the NE of some SCs (red arrows). (B) After 5 h pollen germination, WIT1 was strongly localized at the VN NE and weakly at the SN of SCs (red arrows), while WIP1 is only detectable at the VN NE. Imaging conditions were tuned to view the signal at the SN NE, so that the GFP-WIT1 signal at the VN NE is saturated and looks nuclear. (C) WIT1 is localized normally at the NE of the VN and the SN (red arrows) in sun1-KO sun2-KD (imaged under the same conditions as wild type), but its signal in wip1-1 wip2-1 wip3-1 pollen grains is barely detectable (imaged at 5 times higher laser power and a larger pinhole, which is reflected by the strong autofluorescence from the pollen wall). In (A) - (C), scale bars equal 10 µm.

Figure 3.3 Wit1-1 wit2-1, wip1-1 wip2-1 wip3-1, and wifi pollen conditions. (A) Alexander staining of pollen grains. No obvious dead pollen grains were found in wit1-1 wit2-1, wip1-1 wip2-1 wip3-1, and wifi. (B) Hoechst 33342 DNA staining. The three nuclei were normal in the pollen grains of wit1-1 wit2-1, wip1-1 wip2-1 wip3-1, and wifi. Bars equal 5µm.

Figure 3.4 Nuclear movement in pollen tubes is affected in wit1-1 wit2-1, wip1-1 wip1-1 wip2-1 wip3-1, and wifi. (A) Kymograph of nuclear movement in a wild-type pollen tube with the VN-RFP and the SN-GFP markers. Vertical axis represents time (total=180 min) and horizontal axis represents the distance along the pollen tube growing track (total pollen tube length=223.5 µm). (B) Kymograph of nuclear movement in a wit1-1 wit2-1 pollen tube with the VN-RFP and the SN-GFP markers. Vertical axis represents time (total=180 min) and horizontal axis represents the distance along the pollen tube growing track (total pollen tube length=253.5 µm). (C) Entrance lengths of wild-type and wit1-1 wit2-1 pollen tubes. “**” represents significant statistical difference when compared with wild type (P<0.01, n=40, 2-tailed Student’s t-test). (D) The first nucleus that entered the pollen tubes of wild type and wit1-1 wit2-1. Numbers of each category are shown in each column. “**” represents significant statistical difference when compared with wild type (P<0.01, 2-tailed Fisher’s exact test). (E) Distance between the leading nucleus and the pollen tube tip. The nuclear movement of 10 wild- type pollen tubes and 10 wit1-1 wit2-1 pollen tubes were tracked every minute for 60 min, and the distance was the average of all tracking points. “**” represents significant statistical difference when compared with wild type (P<0.01, n=600, 2-tailed Student’s t- test). (F) Nuclear order in pollen tubes after 5h germination. The example of each category is shown on right. For wip1-1 wip2-1 wip3-1 and wifi, Hoechst 33342 was used to stain DNA for observing nuclear positions, and confocal microscopy was used for the others. Two-tailed Fisher’s exact test was used for statistical analysis and n for each data set is indicated in graph. “**” represents significant statistical difference when compared with wild type (P<0.01), while “o” represents non-significant statistical difference when compared with wild type (P>0.05). Error bars in all histograms represent S.D.

Figure 3.4 Nuclear movement in pollen tubes is affected in wit1-1 wit2-1, wip1-1 wip1-1 wip2-1 wip3-1, and wifi. 88

Figure 3.5 Distance between the leading nucleus and the tip of the growing pollen tube after the nucleus entered the pollen tube. Green color curves, wild-type. Orange color curves, wit1-1 wit2-1. Bold blue curve, the mean value of wild-type curves. Bold red curve, the mean value of wit1-1 wit2-1 curves. Wild-type and wit1-1 wit2-1 mean values are statistically significantly different from each other at every time point (P<0.05, n=10, 2-tailed Student’s t-test). Error bars represent S.D.

Figure 3.6 Nuclear position after 8 h of semi-in-vitro pollen tube germination. Wild-type and wit1-1 wit2-1 pollen grains with the VN-RFP and SN-GFP markers were simultaneously germinated on wild-type stigmas in the same medium. The GFP and RFP signals in the wit1-1 wit2-1 image were enhanced for easy viewing of the nuclear position. Bar=100 µm.

Figure 3.7 Wit1-1 wit2-1 has SC delivery defects. A large number of small ovules was found in wit1-1 wit2-1, wip1-1 wip2-1 wip3-1, and wifi ovaries ~48 h after pollination when compared with wild type. These small ovules were not fertilized, while the large ovules were at the 2-cell to 8-cell stages of embryo development. (B) Wit1-1 wit2-1 had a significantly reduced number of large, fertilized ovules in ovaries ~48 h after pollination, when compared to wild type (“**” represents P<0.01, n=5, 2-tailed Student’s t-test). (C) SC position in small unfertilized ovules. The pie chart shows the percentage of each category of SC positions. The examples are shown on right. Red arrowheads point to the SCs. Scale bar equals 20 µm.

Figure 3.8 Root hair and trichome nuclear shape is regulated by WIT2. (A) Nuclear shape of root hairs and trichomes. Root hair nuclei were imaged by fluorescence microscopy after Hoechst 33342 staining, and trichome nuclei were imaged directly through bright field, except for WIT2pro::GFP-WIT2* in wit1-1 wit2-1 line1 whose nuclei were images by confocal microscopy. (B) Quantified root hair shape. Total nuclear length was used as an index. Eighty nuclei were measured for each line. (C) Quantified trichome nuclear shape. The ratio of width and length was used as an index. Fifty nuclei were measured for each line. In (B) and (C), 2-tailed Student’s t-test was used for analysis—asterisks represent significant statistical difference when compared with wild type (“**” represents P<0.01, “*” represents 0.01

Figure 3.9 WIT2* interact with and WIP1, WIP2, and WIP3. GFP-WIT2* was co-expressed with Myc-WIP1, Myc-WIP2, and Myc-WIP3 in N. benthamiana leaves. As a negative control, NLS-GFP-NES-GFP was co-expressed with Myc-WIP3 in N. benthamiana leaves. Protein extracts were immunoprecipitated with an anti-GFP antibody. The input/IP ratio is 1/9. Arrows denote possible degradation products from GFP-WIT2*. Numbers on left are mol wt in kD.

CHAPTER 4

DORY: an effective algorithm to discover a multitude of novel plant SUN domain-

interacting tail proteins

4.1 Abstract

The nuclear envelope (NE) bridges formed by Sad1/UNC-84 (SUN) proteins and

Klarsicht/ANC-1/Syne-1 Homology (KASH) proteins are essential for nuclear positioning and chromosome movement. While SUN proteins are well-conserved among eukaryotes including plants, no KASH homologs can be found in the plant proteome. The

WPP domain-interacting proteins (WIPs) were recently identified as KASH protein analogs in plants. WIPs have no similarity to known opisthokont KASH proteins, except for a C-terminal transmembrane domain, followed by a short SUN-domain interacting tail

(SIT) domain terminating in a conserved four-amino-acid motif critical for the SUN-WIP interaction. Here, this pattern was used to computationally search for candidate SIT proteins. As a second criterion, we asked for conservation of the rule in most homologs of a protein family. We identified 10 new potential SIT protein families, four of which were verified for their SUN-interaction-dependent NE localization. We further show that

Arabidopsis thaliana AN HEMO 1 is associated with F-actin and is involved in guard cell nuclear central anchorage. This study dramatically expands the number of SIT proteins and implies an independent evolution of SUN-dependent NE-bridging complexes after the opisthokont-plant separation.

4.2 Introduction

At the basic structural level, the nuclear envelope (NE) in all eukaryotes appears alike— double lipid membranes perforated by nuclear pore complexes. However, at the molecular level, the protein complement of the opisthokont NE appears to be largely missing from land plants, while plant-unique NE proteins have been described

(Graumann and Evans, 2010; Gruenbaum et al., 2003; Meier, 2001; Xu et al., 2007; Zhao et al., 2008). This is in stark contrast to the nuclear pore complex, which seems to have been established at the time of the last eukaryotic common ancestor (Neumann et al.,

2010). In light of the fact that open mitosis has likely evolved at least twice (Becker and

Marin, 2009; De Souza and Osmani, 2007; Meier et al., 2008), there may be a much deeper functional and structural divide between opisthokont and land plant NEs.

Opisthokont NE bridging complexes are protein complexes that directly link the nuclear and cytoplasmic environment through the NE. In these complexes, inner nuclear membrane (INM) Sad1/UNC-84 (SUN) proteins interact with outer nuclear membrane

(ONM) Klarsicht/ANC-1/Syne Homology (KASH) proteins through SUN-KASH domain interactions in the perinuclear space (PNS) (Razafsky and Hodzic, 2009; Sosa et al.,

2012; Starr and Fridolfsson, 2010; Zhou et al., 2012b). Also called LINC complexes

(linker of nucleoskeleton and cytoskeleton) (Crisp et al., 2006), they have been identified in multiple organisms and have been shown to play roles in nuclear positioning, nuclear shape, and chromatin-NE interactions, which are essential for a variety of processes including cell division, migration, and polarization (Gundersen and Worman, 2013;

Mellad et al., 2011; Razafsky and Hodzic, 2009; Rothballer and Kutay, 2013; Starr and

Fridolfsson, 2010). Mutations in SUN or KASH proteins lead to a number of developmental abnormalities and are implicated in human diseases (Elhanany-Tamir et al., 2012; Malone et al., 1999; Schreiber and Kennedy, 2013; Starr et al., 2001; Zhang et al., 2009).

SUN proteins are among the few evolutionarily conserved NE proteins also found in plants (Graumann et al., 2010a; Oda and Fukuda, 2011). However, no homologs of the opisthokont KASH protein genes can be identified in the plant genomes. Recently, we have identified Arabidopsis thaliana WPP domain-interacting proteins (AtWIPs) as plant

KASH protein analogs (Zhou et al., 2012a). The Arabidopsis thaliana SUN (AtSUN)-

AtWIP complexes anchor plant Ran GTPase activating protein 1 (RanGAP1) to the NE and are required for the elongated nuclear shape in epidermal cells. WIPs also interact with WPP domain-interacting tail-anchored proteins (WITs) and they synergically anchor

RanGAP1 to the NE (Zhao et al., 2008). Recent evidence shows that SUN-WIP-WIT- myosin XI-I complexes probably exist and regulate elongated nuclear shape and nuclear movement (Tamura et al., 2013), suggesting that the LINC complex is also conserved in plants.

The SUN-WIP complexes are the only known NE bridges in plants. Compared with our understanding of opisthokont KASH proteins, our knowledge of plant SUN-binding NE proteins is limited. Based on detailed knowledge on opisthokont SUN-KASH complex formation (Sosa et al., 2012; Zhou et al., 2012b) and the known conservation of the

KASH domains, we reasoned that a computational approach could be developed to discover unknown KASH protein analogs in plants. Here, we describe this approach and the experimental verification of five candidates from four protein families. One protein family, the AN HEMO (NEMO) family, is conserved across land plants and was studied in detail. We show that the Arabidopsis thaliana NEMO1 is associated with F-actin and play a role in guard cell central nuclear anchorage. The SUN-interacting NE proteins presented here are non-homologous to opisthokont KASH proteins, suggesting that the plant lineage has recruited an independent complement of cytoplasmic proteins into the

SUN-dependent NE bridging complexes.

4.3 Material and Methods

4.3.1 DORY and BLASTP search

DORY was programed in Java. Please refer to Figure 4.1 for details and the source code is available on request. The TAIR10 protein models were downloaded from TAIR

(ftp://ftp.arabidopsis.org/home/tair/Proteins/TAIR10_protein_lists/TAIR10_pep_201012

14). The nr database (ftp://ftp.ncbi.nih.gov/blast/db/FASTA/nr.gz) is constantly updated, which affects the results of DORY and NCBI BLASTP. The one used for DORY here was downloaded on February 19th, 2013. Homologs of newly identified plant putative

SIT (pSIT, see 4.4.1 for details) proteins were obtained by using NCBI BLASTP on June

3rd, 2013. In the result of NEMO homolog search, Cucumis sativus protein GI449441039 shows higher similarity to protein At5G62580.1 (BLASTP Score = 447 bits and E-value

= 1e-125) than to AtNEMO1 (BLASTP Score = 50.8 bits and E-value = 3e-6). Therefore, it was not considered a true homolog of AtNEMO1. For animal KASH protein search, homologs of known animal KASH proteins—Nesprin-1, Nesprin-3, Nesprin-4, Klarsicht,

UNC-83, KASH5, and Lrmp—were obtained by BLASTP. Sequences containing no

Proline in the last four amino acids were submitted to BLASTP and removed if the closest homolog was not a KASH protein. The C4 motif pattern was derived from the final KASH protein pool. All alignments were done by using MAFFT (Katoh and

Standley, 2013) with default settings and the “E-INS-i” strategy, except for the alignment in Figure 4.3G for which the “G-INS-i” strategy was used.

4.3.2 Protein model correction based on the “transmembrane domain (TMD)-pSIT”

architecture

During searching for homologs of newly identified pSIT proteins, we found many proteins share high similarities at the N-termini but lack a pSIT domain. Many of these cases were due to a mis-predicted intron and were then corrected based on the TMD-pSIT architecture. Correction of Selaginella moellendorffii GI302806946 is supported by EST

GI169026300. Correction of Glycine max GI356508173 is supported by EST

GI21889745 and GI6847292. Correction of Glycine max GI356510247 is supported by

EST GI7796284. Correction of Vitis vinifera GI147788255 is supported by EST

GI30321072, GI110420183, and GI33406362. The corrected protein sequences are available on request.

4.3.3 Plant materials

Arabidopsis (Columbia ecotype) were grown at 25⁰C in soil under 16-h light and 8-h dark or on MS (Caisson laboratories) plates with 1% sucrose under constant light. The sun1-KO sun2-KD mutant (SALK_123093 for sun1-KO and SALK_049398 for sun2-

KD) was a gift from Dr. Susan Armstrong (University of Birmingham, UK) and has been reported previously (Zhou et al., 2012a). N. benthamiana plants were grown at 28⁰C in soil under constant light. nemo1-1 (SALK_018239C), nemo1-2 (SALK_143274), nemo2-

1 (CS801355), and nemo2-2 (CS1006876) were ordered from the Arabidopsis Biological

Resource Center. nemo1-3 (GK-485E08-019738) was ordered from GABI-Kat. The

100 primers used for genotyping are listed in Table 4.2. Genotyping of the qrt1-2 allele in nemo2-1 was performed as reported by Francis et al. (2006).

4.3.4 Constructs

CFP-AtSUN2, CFP-AtSUN2ΔN, CFP-AtSUN2ΔCC, CFP-AtSUN2ΔCSUN were described by Graumann et al. (2010a). Myc-Flag-AtSUN1, Myc-Flag-AtSUN2, and GFP-

Myc-AtSUN2ΔNSUN were described previously (Zhou et al., 2012a). PCR-based cloning was used to generate the other constructs and the primers used are listed in Table 4.1.

Lifeact was amplified by PCR using self-annealing primer pair without template and cloned to pENTR/D-TOPO (Invitrogen). RT-PCR was used to amplify AtNEMO1,

AtNEMO2, AtNEMOL, and MsNEME1 from cDNA. AtNEMOS was amplified directly from wild-type genomic DNA by PCR. NLS-GFP-NES was amplified by overlapping

PCR using vector pK7WGF2 as a template. All PCR products were cloned into the pENTR/D-TOPO vector and confirmed by sequencing. These clones were then used as templates for amplifying AtNEMO1ΔTVPT, AtNEMO1XT, AtNEMO11-308,

AtNEMO2ΔLVPT, AtNEMO2XT, AtNEMO21-309, AtNEMO2∆KASH, AtNEMOLΔPLPT,

AtNEMOLXT, AtNEMOSΔLVPT, and MsNEME1ΔLVPT by PCR. PCR produces were cloned into pENTR/D-TOPO vector and confirmed by sequencing. The coding sequences cloned in pENTR/D-TOPO were then moved to destination vectors described by (Karimi et al., 2002) by LR reaction (Invitrogen) to obtain N-terminal-GFP-tagged protein constructs: Lifeact was cloned into pH7WGR2; AtNEMO1, AtNEMO1ΔTVPT,

AtNEMO1XT, AtNEMO11-308, AtNEMO2, AtNEMO2ΔLVPT, AtNEMO2XT, AtNEMO21-

101

309, AtNEMO2∆KASH, AtNEMOLΔPLPT, AtNEMOLXT, AtNEMOS, AtNEMOSΔLVPT,

MsNEME1ΔLVPT, and NLS-GFP-NES were cloned into the pK7WGF2; AtNEMOL and

MsNEME1 were cloned into pH7WGF2. Flag-AtSUN1dMut and Myc-AtSUN2dMut were obtained by using the QuikChange® Site-Directed Mutagenesis Kit and the pENTR/D-

TOPO vector containing the Flag-AtSUN1 or the Myc-AtSUN2 coding sequence described by Zhou et al. (2012a) was used as a template. After sequencing, Flag-

AtSUN1dMut and Myc-AtSUN2dMut were cloned into pGWB21 (Nakagawa et al., 2007) to obtain Myc-Flag-AtSUN1dMut and Myc-AtSUN2dMut, respectively.

AtNEMO1 and AtNEMO2 promoters were amplified from Arabidopsis genomic DNA (~

2kb), digested with AscI, and linked to the AscI/PmlI-digested pHOAG (see CHAPTER

3) to obtain pHAtNEMO1proAG and pHAtNEMO2proAG, respectively. AtNEMO1 CDS was moved from the pENTR/D-TOPO to pHAtNEMO1proAG to obtain

AtNEMO1pro::GFP-AtNEMO1 construct. AtNEMO2 CDS was moved from the pENTR/D-TOPO to pHAtNEMO2proAG to obtain AtNEMO2pro::GFP-AtNEMO1 construct.

4.3.5 Agrobacterium transformation, N. benthamiana transient expression, and

Arabidopsis stable transformation

Agrobacterium tumefaciens stain ABI was transformed with the corresponding constructs by triparental-mating (Wise et al., 2006). Agrobacterium cultures containing plasmids expressing the proteins of interest were co-infiltrated transiently into N. benthamiana

102 leaves as described previously (Sparkes et al., 2006). Transgenic Arabidopsis plants were obtained by Agrobacterium-mediated floral dip (Clough and Bent, 1998).

4.3.6 Co-immunoprecipitation experiments

Plants were grown for 3 days after infiltration at 28⁰C under constant light. Leaves were collected, ground in liquid nitrogen into powders, and Co-IP experiments were performed at 4⁰C. RIPA buffer containing 50 mM Tris-HCl, pH7.5, 150 mM NaCl, 0.1% SDS,

0.5% NaDeoxycholate, 1% NP-40, 1 mM PMSF, and 1% protease inhibitor cocktail

(Sigma-Aldrich) was used. Protein complexes were immunoprecipitated from extracts by an anti-GFP antibody (ab290, Abcam Cambridge) bound to protein A-sepharose beads

(GE Healthcare). After three times wash using RIPA buffer, the immunoprecipitates and the input samples were separated by 8% or 10% PAGE, transferred to PVDF membrane

(Bio-Rad), and detected with an anti-GFP (1:2000, 632569, Clontech) or anti-Myc

(1:1000, M5546, Sigma-Aldrich) antibody. Please see 2.3.4CHAPTER 2 for details.

4.3.7 Confocal microscopy

Seven- to 10-day old Arabidopsis seedlings were imaged using a Nikon Eclipse C90i confocal microscope with small or medium pinhole and gain setting of 7.0 to 7.5. The

488nm laser was set at 40% power for imaging sun1-KO sun2-KD transgenic plants whose transgene expression was low in lines, while the other transgenic lines and N. benthamiana leaves were imaged using 10%-20% laser power. All images were taken at

103 room temperature with Nikon Plan Apo VC 60x H lens (numerical aperture 1.4). The transmitted light detector was turned on to collect transmitted light signal simultaneously.

4.3.8 Nuclear position measurement in guard cells

Leaves of 4-6 weeks Arabidopsis plants were used for the nuclear position measurement in guard cells. To open stomata, newly fully expanded leaves were detached, put on a piece of wet filter paper in a petri dish, and irradiated with 100 µmol/m2/s blue light for 3 h. The lower epidermis was then peeled off and fixed and stained in 4% paraformaldehyde PBS buffer (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM

KH2PO4, pH7.4) containing 4 µM Hoechst 33342 for at least 20 min. After this fixation and staining, the leaves were imaged using a Nikon DS-Qi1Mc digital camera. Leaves without blue-light treatment were fixed, stained, and imaged in the same way, except for the AtNEMO1pro::GFP-AtNEMO1 transformed nemo1-1 lines which were imaged directly on a Nikon C90i confocal microscope using the fluorescence from GFP-

AtNEMO1.

Nikon NIS-elements software was used for the nuclear position measurement. An ellipse was first rendered on a pair of guard cells using the 5-point ellipse tool. The major axis of the ellipse was aligned to the common boundary of the two guard cells. If the major axis was equal to or longer than 8 µm, the angles between the minor axis and the middle of the nuclei were measured using the free angle tool.

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4.4 Results

4.4.1 Identify Arabidopsis thaliana SUN-interacting protein candidates

The SUN-domain interacting tail (SIT) of opisthokont KASH proteins and plant WIPs terminate in a four-amino-acid motif (C4 motif). The SIT domain, especially the C4 motif, is critical for interacting with the SUN domain and for the NE localization

(Ketema et al., 2007; Morimoto et al., 2012; Padmakumar et al., 2005; Sosa et al., 2012;

Zhou et al., 2012a). However, the SIT of plant WIPs and of animal KASH proteins shares little amino acid similarity. Even among plant WIPs, the composition of the SIT domain varies significantly, except for the conserved C4 motif, and in particular, the terminal

“PT” motif (Figure 4.1A). Thus, we define a protein sequence as a putative SIT (pSIT) when it fulfills all of the following: (1) it is immediately C-terminal of a TMD; (2) its length is between 9-40 amino acids (based on animal KASH proteins and plant WIPs);

(3) it is the C-terminus of a protein and terminates in four amino acids with a given amino acid pattern. A Java program named “DORY” (DOes it RobustlY) was developed to search for pSIT-containing proteins according to these rules (see flow chart in Figure

4.1B). In addition, we argued that if a pSIT is present in most homologs of a protein family, it is more likely to be a bona fide SIT domain. Therefore, BLASTP was used to obtain homologs of a positive output from DORY in the NCBI non-redundant (nr) database (in this study, proteins with E-value<=0.0001 are considered as homologs).

To develop the C4 motif pattern of plant SIT for DORY, we searched the Arabidopsis thaliana TAIR10 protein models using DORY with default settings and the C4 motif

105 pattern “XXPT” (X represents any amino acid). Five new protein models were identified:

At1G54385.1, At3G03970.1, At3G06600.1, At4G24950.1, and At3G47410.1 (alternative gene models resulting in the same protein sequences were discarded). At1G54385.1 and

At3G03970.1 are paralogs in Arabidopsis thaliana and were named AtNEMO1 and

AtNEMO2, respectively (see below for full gene names). Running BLASTP with

AtNEMO1 revealed that NEMO homologs exist across land plants (Figure 4.2A). All

AtNEMO1 homologs were submitted to the TMD prediction program “Phobius”

(http://phobius.sbc.su.se/) (Kall et al., 2007) for verifying the existence of a TMD.

Although Physcomitrella patens GI168052080 was not predicted to possess a TMD, it contains a ~20 amino acid hydrophobic fragment at the conserved position, likely a false- negative prediction by Phobius. Among the 55 homologs (E <= 0.0001) found, 52 terminate in a pSIT domain, suggesting that this protein family may be a bona fide SIT protein family.

At3G06600.1 (AtNEMOL, see below for naming) has close homologs in Arabidopsis lyrata and in the closely related species Capsella rubella. BLASTP also discovered several homologous proteins in other plant species with relatively high E-values (0.003-

0.064), indicating that there are distantly related proteins. Using PSI-BLAST with an E- value threshold of 0.005, the number of homologs converged at the fifth iteration. As shown in Figure 4.2B, 8 of the 11 homologs terminate in a pSIT domain. At4G24950.1

(AtNEMOS, see below for naming) has only one homolog each in Arabidopsis lyrata and

Capsella rubella. All three homologs possess a pSIT domain. At3G47410.1 has one

106 homolog in Arabidopsis thaliana, two in Arabidopsis lyrata, and one in Capsella rubella.

Two Arabidopsis homologs do not possess a pSIT (data not shown). It was therefore considered false-positive.

Analyzing the putative C4 motif of homologs of WIP, NEMO, NEMOL, and NEMOS revealed a new pattern of “[DTVAMPLIFY][VAPIL]PT” (brackets indicate alternative amino acid residues at the respective position). This pattern was used to search the nr database for pSIT proteins in plants, using DORY with default settings. Seven new protein families were identified (see Figure 4.2D and Figure 4.3 A-G). The ORFs of

Oryza sativa GI242067046 and GI125591230, as well as Triticum urartu GI473759431

(Figure 4.3 A and B) continue beyond the “PT” motif and were not considered to possess a pSIT domain. Nonetheless, because the majority of these protein families contain a pSIT, they were included as putative plant SIT proteins. The protein family shown in

Figure 4.2D was identified only in Medicago truncatula and was subsequently cloned in

Medicago sativa, too. The proteins shown in Figure 4.3 A and B were only found in

Poaceae.

The search with these criteria also identified a mammalian protein family, transmembrane

191C (T191C), which has a conserved TMD followed by a short sequence (~40 amino acid) terminating in a conserved “LLP[AST]” motif (Figure 4.3G; see below).

4.4.2 Verify the subcellular localization of SUN-binding protein candidates

AtNEMO1, AtNEMO2, AtNEMOL, AtNEMOS, and the Medicago sativa homolog of

Medicago truncatula GI357448889 (MsNEME1) were chosen to determine their 107 subcellular localization. Transgenic Arabidopsis lines were generated that express N- terminally GFP-tagged proteins under the control of the Cauliflower Mosaic Virus 35S

(35S) promoter. Root tips cells of 7-10 day old seedlings were imaged by confocal microscopy, and at least three lines for each construct were analyzed. As shown in Figure

4.4A, all five fusion proteins were associated with the NE. Based on the localization,

At1G54385 and At3G03970 were named AN HEMO 1 and 2, respectively (NEMO; AN,

German, “at”; HEMO, Chinese, “nuclear envelope”). Accordingly, we named

At3G06600 AN HEMO LITTLE (NEMOL) and At4G24950 AN HEMO SMALL

(NEMOS), based on the shortness of their ORFs. GI357448889 and GI357488369 were named AN HEMO IN MEDICAGO 1 (NEME1) and NEME2, respectively.

4.4.3 AtNEMO1, AtNEMO2, AtNEMOL, AtNEMOS, and MsNEME1 interact with

AtSUN1 and AtSUN2

Next, we tested the ability of AtNEMO1, AtNEMO2, AtNEMOL, AtNEMOS, and

MsNEME1 to interact with AtSUN1 and AtSUN2. Pairs of tagged proteins were transiently expressed under the control of the 35S promoter in Nicotiana benthamiana leaves and co-immunoprecipitation (IP) assays were performed. To determine the importance of the putative C4 motif, deletions of last four amino acids were introduced into all five tested proteins and designated as AtNEMO1ΔTVPT, AtNEMO2ΔLVPT,

AtNEMOLΔPLPT, AtNEMOSΔLVPT, and MsNEME1ΔLVPT, respectively (Figure

4.5A and Figure 4.6A). To test whether pSIT is essential for SUN protein interaction, the pSIT of AtNEMO1, AtNEMO2, and AtNEMOL was replaced with

108

“RFYTKSAEAAAAA” (AtNEMO1XT), “RFYTKSAEAAAA” (AtNEMO2XT), and

RFYTKSAEAAAAAA (AtNEMOLXT), respectively (Figure 4.5A). The underlined sequence is the PNS domain of the ER tail-anchored protein cytochrome b5c from

Aleurites fordii (Hwang et al., 2004), lengthened with alanine residues to the size of the respective pSIT.

GFP-AtNEMO1, GFP-AtNEMO1ΔTVPT, GFP-AtNEMO1XT, GFP-AtNEMO2, GFP-

AtNEMO2ΔLVPT, or GFP-AtNEMO2XT was transiently co-expressed with N- terminally Myc-Flag-tagged AtSUN1 (Myc-Flag-AtSUN1). IP assays were performed with an anti-GFP antibody. As shown in Figure 4.5C, Myc-Flag-AtSUN1 was strongly co-immunoprecipitated with GFP-AtNEMO1 or GFP-AtNEMO2 but barely with GFP-

AtNEMO1ΔTVPT, GFP-AtNEMO2ΔLVPT, GFP-AtNEMO1XT, or GFP-

AtNEMO2XT. A similar co-IP procedure was performed to test the interaction of Myc-

AtSUN2 with AtNEMO1, AtNEMO2, and their respective mutants. As shown in Figure

4.5D, Myc-AtSUN2 was strongly co-immunoprecipitated with GFP-AtNEMO1 or GFP-

AtNEMO2, weakly with GFP-AtNEMO1ΔTVPT, and barely with GFP-

AtNEMO2ΔLVPT, GFP-AtNEMO1XT, or GFP-AtNEMO2XT. These data indicate that the pSIT domain and the terminal C4 motif are important for AtSUN1 and AtSUN2 binding.

To test whether the SUN domain of AtSUN1 and AtSUN2 is necessary for this interaction, co-IP assays were carried out with AtSUN2 deletion mutants (Figure 4.5B) and AtNEMO1. Myc-AtNEMO1 was co-expressed with CFP-AtSUN2, CFP-AtSUN2ΔN

109

(deletion of the N-terminal 106 aa), CFP-AtSUN2ΔCC (deletion of the coiled-coil domain, aa 205-225), CFP-AtSUN2ΔCSUN (deletion of the C-terminal 146 aa of the

SUN domain) and GFP-Myc-AtSUN2ΔNSUN (deletion of the N-terminal 84 aa of the

SUN domain). As a negative control, the unrelated GFP-NLS-GFP-NES (GFP fused with a nuclear localization signal and a nuclear export signal) was used. After IP using an anti-

GFP antibody, co-immunoprecipitated Myc-AtNEMO1 was detected by an anti-Myc antibody. As shown in Figure 4.5E, only the two SUN-domain deletions were unable to co-immunoprecipitate Myc-AtNEMO1.

To further analyze SUN-domain specificity of the protein-protein interactions, we introduced two point mutations in the SUN domain of AtSUN1 and AtSUN2. Based on the amino acid sequence alignment of AtSUN1, AtSUN2, and Homo sapiens SUN2 (see

Figure 1.2 and Figure 1.3 in CHAPTER 1) (Zhou and Meier, 2013), two conserved residues in the KASH-binding pocket were chosen and mutated in AtSUN1 (H439A and

Y443F) and AtSUN2 (H434A and Y438F) (illustrated in Figure 4.5B). The mutated proteins were named AtSUN1dMut and AtSUN2dMut, respectively. After N. benthamiana co-expression and co-IP, the ability of GFP-AtSUN1dMut and GFP-

AtSUN2dMut to bind Myc-AtNEMO1 or Myc-AtNEMO2 was determined. As shown in

Figure 4.5 F and G, in contrast to wild-type SUN proteins, both mutated proteins barely interacted with either AtNEMO1 or AtNEMO2, confirming that the KASH-binding pocket within the SUN domain is required for interaction.

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Figure 4.6 shows the comparable co-IP assays performed for GFP-tagged AtNEMOL,

AtNEMOS, and MsNEME1, and their respective mutants, tested for interactions with

Myc-tagged AtSUN1, AtSUN1dMut, AtSUN2, and AtSUN2dMut. IPs were performed with an anti-GFP antibody, and co-immunoprecipitated proteins were detected with an anti-Myc antibody. Figure 4.6A shows schematic representation of AtNEMOL,

AtNEMOS, MsNEME1, and their pSIT domain mutants. Figure 4.6 B and C show that

AtNEMOL strongly binds to AtSUN1 and AtSUN2, but that either the deletion of the C- terminal 4 amino acids or replacement of the pSIT tail or mutating the KASH-binding pocket of AtSUN1 and AtSUN2 disrupts this binding. Similar results were obtained for

AtNEMOS (Figure 4.6 D and E) and for MsNEME1 (Figure 4.6 F and G).

4.4.4 The NE Localization of AtNEMO1, AtNEMO2, AtNEMOL, AtNEMOS, and

MsNEME1 depends on AtSUN

Losing interaction with a SUN protein leads to reduced NE localization of Nesprin-2

(Crisp et al., 2006; Padmakumar et al., 2005), Nesprin-3 (Ketema et al., 2007), Nesprin-4

(Roux et al., 2009), KASH5 (Morimoto et al., 2012), and WIP1 (Zhou et al., 2012a). To determine whether this is true for GFP-AtNEMO1ΔTVPT, GFP-AtNEMO2ΔLVPT,

GFP-AtNEMOLΔPLPT, GFP-AtNEMOSΔLVPT, GFP-MsNEME1ΔLVPT, they were stably expressed in Arabidopsis under the control of the 35S promoter. Figure 4.4B shows that all five GFP fusion proteins were only weakly associated with the nuclear periphery and were abundantly found in the cytoplasm and/or associated with the plasma membrane and components of the endomembrane system. This demonstrates that the C4

111 motif is required for efficient association of all five proteins with the NE. Finally, all five

GFP-fusion proteins were expressed in the sun1-KO sun2-KD double mutant (Zhou et al.,

2012a). Figure 4.4C shows that this led to a strong loss of NE localization, comparable to the C4 motif deletion.

4.4.5 The armadillo-repeat (ARM) domain of AtNEMO1 is associated with F-actin

Since the NEMO family is conserved across the land plants, we investigated AtNEMO1 and AtNEMO2 in detail. InterProScan (http://www.ebi.ac.uk/Tools/pfa/iprscan/) predicts

ARM folds in the N-termini of AtNEMO1 and AtNEMO2 (aa 3-286 and aa 17-289, respectively). We tagged the N-terminal 308AA of AtNEMO1 with GFP (GFP-

AtNEMO11-308, containing the ARM domain) and transiently expressed this fusion protein in N. benthamiana leaves under 35S promoter. Interestingly, GFP-AtNEMO11-308 was localized to cytoplasmic fiber-like structures, most likely F-actin fibers. We then co- expressed GFP-AtNEMO11-308 with an F-actin marker, RFP-Lifeact (Riedl et al., 2008), or a microtubule marker, MAP4-RFP, in N. benthamiana leaves. As shown in Figure

4.7A, GFP-AtNEMO11-308 was co-localized with RFP-Lifeact but not with MAP4-RFP.

The co-localization of GFP-AtNEMO11-308 with RFP-Lifeact was also observed in stably transformed Arabidopsis (Figure 4.7A). However, the N-terminal 309AA of AtNEMO2

(AtNEMO21-309, containing the ARM domain) was not localized to any fiber-like structures in N. benthamiana leaves (Figure 4.7A), neither was the whole AtNEMO2 N- terminus without the TMD-SIT domain (AtNEMO21-521, Figure 4.7B), suggesting that the F-actin association property is specific to AtNEMO1.

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4.4.6 AtNEMO1 has a specific expression and localization pattern

To analyze the expression profiles of AtNEMO1 and AtNEMO2, GFP-GUS driven by

AtNEMO1 promoter (AtNEMO1pro:: GFP-GUS) and AtNEMO2 promoter

(AtNEMO2pro:: GFP-GUS) were stably transformed to wild-type Arabidopsis. Three transformed lines for each construct were examined and representative images are shown in Figure 4.8A. Both AtNEMO1 and AtNEMO2 were expressed in roots and leaves. For detailed gene expression and protein localization information, we imaged the N- terminally GFP-tagged AtNEMO1 and AtNEMO2 driven by their own promoter

(AtNEMO1pro::GFP-AtNEMO1 and AtNEMO2pro::GFP-AtNEMO2) in stably transformed wild-type Arabidopsis, respectively. In leaves, AtNEMO1 was exclusively expressed in guard cells and guard cell mother cells, while AtNEMO2 was expressed in epidermal cells, mesophyll cells, trichomes, but weakly in guard cells and guard cell mother cells (Figure 4.8B, C, and D). Interestingly, AtNEMO1 showed a spotted fiber- like pattern at the NE and in the cytoplasm of guard cells and guard cell mother cells

(Figure 4.8C). In root cells, the fiber-like pattern of AtNEMO1 was also visible at the NE

(Figure 4.8E). However, AtNEMO2 showed no fiber-like localization pattern in either leaves or roots (Figure 4.8D and E).

Since the N-terminus of AtNEMO1 is associated with F-actin, the fiber-like structures at the NE of root and guard cells probably depend on F-actin fibers. After treatment with 10

µM Latrunculin B (LatB) for 1 h, the AtNEMO1 NE fiber-like structures disappeared

(Figure 4.8F). In contrast, in the mock treatments, these structures were barely affected,

113 supporting our hypothesis that AtNEMO1 forms F-actin dependent fibers at the NE

(Figure 4.8F).

4.4.7 AtNEMO1 is required for proper nuclear anchorage in guard cells

To dissect the function of AtNEMO1 and AtNEMO2, three AtNEMO1 T-DNA insertion mutants (nemo1-1, nemo1-2, and nemo1-3) and two AtNEMO2 T-DNA insertion mutants

(nemo2-1 and nemo2-2) were isolated. Further analysis showed that at least one downstream gene is missing in nemo1-2, which was therefore discarded. Nemo2-1 carries the homozygous qrt1-2 allele (Francis et al., 2006) which was segregated out in the nemo1-1 nemo2-1 double mutant. All T-DNA insertion sites were confirmed by sequencing (Figure 4.9A). No whole-plant phenotype was observed in nemo1-1, nemo1-

3, nemo2-2, nemo1-1 nemo2-1, nemo1-1 nemo2-2, or nemo1-3 nemo2-2 mutants (Figure

4.10).

Since AtNEMO1 is specifically expressed in guard cells and guard cell mother cells in leaves, we examined the nuclear position in guard cells. Epidermis was peeled and immediately fixed in 4% paraformaldehyde containing 4 µM Hoechst 33342. An ellipse was rendered on a pair of guard cells and the acute angle between the minor axis of the ellipse to the center of the nucleus was used to quantify the position of the nucleus

(Figure 4.9B). To exclude immature guard cells, only guard cell pairs with a major axis of the rendered ellipse longer than 8 µm were measured. To avoid possible stoma aperture effects on nuclear position, we irradiated the leaves with 100 µmol/m2/s blue light to open stomata before fixation. Eighty-six pairs of guard cells were measured for

114 each line and Figure 4.9C shows the distribution of guard cell nuclear position. In wild type and nemo2-2, the guard cell nuclei predominantly localized at the center of the cell.

However, in mutants containing homozygous nemo1-1or nemo1-3 alleles, the position of the nuclei was less well confined, and more often skewed towards a greater distance from the center of a guard cell. We then measured the nuclear position without blue light illumination. As shown in Figure 4.9D, similar results were obtained, suggesting that stoma aperture has little effect on nuclear positioning in guard cells. To accurately compare the difference between wild type and mutants, the number of nuclei below or above the wild-type angle median (11°) was counted, and one-tailed Fisher’s exact test was employed to test the significant difference among different lines. As shown in Figure

4.9F, blue light illumination did not lead to significant difference of the nuclear positions of wild type (P>0.05, n=172), and neither did blue light have effects on nemo1-1 and nemo1-1 nemo2-2 mutants (P>0.05, n=172). The nuclear position of mutants containing either a homozygous nemo1-1 or a homozygous nemo1-3 allele are significantly different from that of wild type (P<0.05, n=172), while nemo2-2 shows no significant difference to wild type (P>0.05, n=172). We then measured the nuclear position of

AtNEMO1pro::GFP-AtNEMO1-transformed nemo1-1. Three lines were analyzed, and in all cases AtNEMO1pro::GFP-AtNEMO1 rescued the nuclear position of nemo1-1, as shown in Figure 4.9 E and F (compared with wild type, P>0.05; compared with nemo1-1,

P<0.01; n=172).

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Guard cell nuclear position was also affected in sun1-KO sun2-KD. As shown in Figure

4.9 E and F, the nuclear position of sun1-KO sun2-KD is similar to that of nemo1-1

(P>0.05, n=172) and is affected when compared with wild type (P<0.05, n=172). These data suggested that the AtSUN-AtNEMO1 complex is involved in the central nuclear anchorage in guard cells.

4.5 Discussion

4.5.1 DORY as a useful tool to identify potential SIT proteins

In this study, DORY predicted 10 new plant SIT protein families. Five members from 4 predicted families were verified to be bona fide plant SIT proteins. The success suggested that DORY had robust and efficacious predictive power. Using the C4 motif pattern

“PPPX” and the nr database, DORY predicted animal coiled-coil domain-containing protein 155 (Ccdc155) and lymphoid-restricted membrane protein (Lrmp) as potential

KASH proteins. Ccdc155 was recently published as KASH5 (Morimoto et al., 2012), and

Lrmp was also suggested to be a KASH protein (Lindeman and Pelegri, 2012). Thus, we used DORY to deep search for unrecognized animal pSIT proteins. Homologs of known animal KASH proteins obtained by BLASTP, and the C4 motif patterns

“[PATHQL]PP[QTVFILM]”, or very rarely “PLPV”, “PSPT”, or “PPKA” were derived.

The pattern “[PATHQL]PP[QTVFILM]” was used for searching for animal KASH proteins in the nr database using DORY with “Maximum SIT Length” set to 60. Five new potential KASH proteins were identified. Their C-termini were aligned with the C- termini of T191C and known animal KASH proteins in Figure 4.3G. Loa loa 116

GI312089182, Wuchereria bancrofti GI402593023, and Brugia malayi GI170594686 are homologs and can only be found in these filarial nematodes. Branchiostoma floridae

GI260805382 is species-specific.

4.5.2 Diversity of plant SIT proteins and opisthokont KASH proteins

The SUN domain is well-conserved among eukaryotes (Starr and Fridolfsson, 2010).

AtSUN1dMut and AtSUN2dMut point mutations were informed by the corresponding

KASH-binding pocket of mammalian SUN2 and abolish plant SIT-binding ability. This indicates that the plant SUN domain and mammalian SUN domain share both sequence and structural similarities. In contrast, there is little to no sequence similarity between the plant SIT domains and opisthokont SIT domains. The plant SIT domains are much shorter (approximately 9-16 amino acids) than those of most opisthokont KASH proteins

(around 30 amino acids). However, the crystal structure of the SUN trimer indicated that the length of the entire luminal domain of Homo sapiens SUN2 was predicted to span the

NE lumen, indicating that this may be also the case for plant SUN proteins and the shorter plant SIT domains should be sufficient to reach the SUN domain (Sosa et al.,

2012).

The KASH protein families revealed here differ vastly in terms of their conservation within the land plant lineage. NEMO is conserved across land plants, including non- vascular plants. This suggests an ancient appearance and makes NEMO family members exciting candidates to probe into an early, conserved function of plant NE bridging complexes. In contrast, WIP is conserved in flowering plants while NEMOL is only

117 found in dicots, indicating a much later appearance of these SUN-binding partners. Other proteins, including NEMOS and NEME are present either in only a few closely related or even a single species, suggesting rather specific functions.

A similar pattern can be derived by comparing prevalence among opisthokont KASH proteins. Homologs of ANC-1, MSP-300, Nesprin-1, and Nesprin-2 are widely conserved in animals (Starr and Fridolfsson, 2010), Klarsicht homologs can only be found in insects

(our BLASTP search, proteins with e-value<=0.0001 were considered as homologs), and

KDP-1 homologs are specific to several nematodes (McGee et al., 2009). No close homologs of S. pombe Kms1 and Kms2 can be found in other species (our BLASTP search, proteins with e-value<=0.0001 were considered as homologs). This “cross kingdom” and “within kingdom” differentiation of SIT/KASH proteins implies that they have emerged after the rise of SUN proteins and that they evolved rapidly to accommodate various functionalities. Given the diverse sequences of the N-terminal domains of plant SIT proteins, detailed future study will be required to understand their functions in the land plant lineage.

4.5.3 Function of AtNEMO1 in guard cell nuclear anchorage

The specificity of animal LINC complexes is determined by the cytoplasmic domains of the KASH proteins. None of the proteins identified here has similarity to the cytoplasmic domains of opisthokont KASH proteins. The N-termini of AtNEMO1 and AtNEMO2 contain ARM domains. Aside from this possible protein-protein interaction domain, no signature or structural elements could be assigned to the cytoplasmic domains of the

118 proteins identified here. The ARM domain of AtNEMO1 is co-localized with F-actin filaments and AtNEMO1 forms LatB-sensitive fiber structures at the NE, suggesting that

AtNEMO1 links SUN proteins to F-actin. The fiber pattern at the NE mimics the actin- associated nuclear (TAN) lines formed by SUN2 and Nesprin-2 found in mammalian fibroblasts (Luxton et al., 2010). TAN lines transfer forces from F-actin to the nucleus, which mediates the backward movement of the nucleus during fibroblast polarization

(Luxton et al., 2010). One significant difference between the AtNEMO1 NE fibers and the TAN lines is that the former are interwoven (Figure 4.8C and E) but the latter are parallel. This could indicate that directional moving of TAN lines requires the fibers to be parallel but interwoven fibers are more stable for the nuclear anchorage function of

AtNEMO1.

Unlike AtNEMO1, mammalian Nesprin-2 and its homologs interact with F-actin through their N-terminal α-actinin-type actin binding domain (Zhang et al., 2002). For ARM- containing proteins, there are two known examples that are associated with F-actin—

Arabidopsis ARMADILLO REPEAT ONLY1 (ARO1) (Gebert et al., 2008) and

Saccharomyces cerevisiae Vac8p (Wang et al., 1998).ARO1 is specifically expressed in pollen vegetative and egg cells and is involved in pollen tube growth and actin organization (Gebert et al., 2008). Vac8p is localized to the vacuole membrane and interacts with Vac17p, an adaptor that binds a myosin V, Myo2p (Wang et al., 1998).

This triple complex regulates vacuole movement and inheritance in yeast (Tang et al.,

2003; Wang et al., 1998). Since AtNEMO1 is predominantly expressed in guard cells, a

119 cell type containing large vacuoles, it is worth further investigation to determine whether it also has a vacuole-related function.

4.5.4 The LINC function of plant SUN-KASH NE bridges

The plant SUN-WIP NE bridges were recently linked to the actin cytoskeleton-network.

Arabidopsis thaliana WPP domain-interacting tail-anchored protein 1 (AtWIT1) and

AtWIT2 interacting with AtWIP1, 2, 3 recruit myosin XI-I to the NE (Tamura et al.,

2013). Loss of myosin XI-I or and AtWIT1/2 lead to restricted nuclear movement and altered nuclear morphology in root epidermal cells. Although the details of the potential

SUN-WIP-WIT-myoxin XI-I complexes require further investigation (see CHAPTER 3), this finding suggests that LINC complexes also exist in plants. The F-actin association and nuclear anchorage function of AtNEMO1 identified in this study provide an additional example of a plant LINC complex.

Many animal F-actin binding KASH proteins, like Nesprin-1, Nesprin-2, and MSP300, are giant proteins containing spectrin repeats (Padmakumar et al., 2004; Zhang et al.,

2002; Zhen et al., 2002). Spectrin repeats’ elastic property is proposed to be important for buffering mechanical stress (Autore et al., 2013; Djinovic-Carugo et al., 2002; Grum et al., 1999; Lenne et al., 2000), and they also form docking sites for protein-protein interactions (Djinovic-Carugo et al., 2002). No KASH proteins of this size have been found in plants so far. There are several possible explanations: 1) the rigid wall of plant cell makes it unnecessary for protecting the nucleus from mechanical stress; 2) the plant nucleus is elastic by itself (the Arabidopsis root nuclei are very dynamic in shape) and

120 does not require mechanical buffering; 3) the elasticity and protein-docking function is fulfilled by other domains in plants. It is noteworthy that the N-termini of AtWIPs and

AtWITs contain interacting coiled-coil domains, elastic structures found in a variety of proteins involved in mechanical and structural tasks (Rose and Meier, 2004). It is also known that the coiled-coiled domain of AtWIT1 interacts with RanGAP and myosin XI-

I, serving as a docking site. Based on these data, it is reasonable to speculate that the coiled-coil domain of AtWIPs and AtWITs is a plant alternative of spectrin repeats in mammalian KASH protein. For AtNEMO1, it is worth further investigating whether it is associated with F-actin through an “elastic” adaptor.

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Figure 4.1 Development of DORY Algorithm. (A) Alignment of WIP homologs. Full- length proteins were used for alignment (MAFFT with “E-INS-i strategy”) and only the C-termini are shown. Species name abbreviations: Aly, Arabidopsis lyrata; Bdi, Brachypodium distachyon; Csa, Cucumis sativus; Gma, Glycine max; Hvu, Hordeum vulgare; Mtr, Medicago truncatula; Osa, Oryza sativa; Ptr, Populus trichocarpa; Rco, Ricinus communis; Sbi, Sorghum bicolor; Tae, Triticum aestivum; Vvi, Vitis vinifera; Zma, Zea mays. Numbers following abbreviations are GI numbers. All other symbols are as in Figure 4.1. ClustalX color was assigned to the alignment. (B) Simplified DORY program flow chart. DORY consists of two filters—the SITtailFilter and the HomologyFilter. In the SITtailFilter, the input protein sequence is scanned over a 20 amino-acid frame (adjustable in “TMD Frame Length”). The total hydrophobic value of the 20 amino acid in this frame is calculated using the Kyte-Doolittle hydrophobicity scale. If the hydrophobicity is greater than or equal to 32 (adjustable in “TMD Hydrophobic Threshold”), this frame is considered a TMD. Proteins that have more than one TMD are discarded, since known KASH proteins have only one TMD. The C- terminal fragment of the TMD is subsequently checked for fulfillment of a pSIT, and the positives are saved in a file named “SITtailFilterResult.txt”. This process is called “SIT search”. To facilitate the analysis of the positive proteins, the HomologyFilter reads proteins in the “SITtailFilterResult.txt” file, categorizes them into homologous groups, and saves each group in a numbered file. This step is called “Homology grouping”. Homology is calculated using the Smith-Waterman algorithm with the BLOSUM62 matrix, the gap open penalty equal to -11, and the gap extension penalty equal to -1. The Smith-Waterman Java code was downloaded from Yang Zhang’s lab (University of Michigan http://zhanglab.ccmb.med.umich.edu/NW-align/SWalign.java.tar.gz) with modification to fit DORY. E-value was calculated by the equation Kmne−λS. S is the score of an alignment. K and λ are Karlin-Altschul parameters whose values are obtained from the BLAST source code. Parameters m and n are the effective lengths of the query sequence and database, respectively, which are calculated by a modified “BLAST_ComputeLengthAdjustment” function from the BLAST source code. E- value<0.0001 is the default setting (adjustable in “E-value Cutoff”) for considering two protein sequences as homologs. DORY also labels whether an output group contain homologs more than a “Homolog Cutoff” value (the default value is arbitrarily set to 4). If not, this output file is labeled with the prefix “belowHomoCutOff”. To make the prediction of the TMD more accurate, positive proteins were manually submitted to the Phobius program for verifying the existence of a TMD.

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Figure 4.1 Development of DORY Algorithm 123

Figure 4.2 Amino acid sequence alignment of the C-terminal domains of predicted plant SIT proteins. Amino acid sequence alignment of NEMO homologs (A), NEMOL homologs (B), NEMOS homologs (C), and NEME homologs (D). Full-length protein sequences were used for the alignment, and only the C-termini are shown here. Aly, Arabidopsis lyrata; Ata, Aegilops tauschii; Bdi, Brachypodium distachyon; Cru, Capsella rubella; Csa, Cucumis sativus; Fve, Fragaria vesca; Gma, Glycine max; Hvu, Hordeum vulgare; Mtr, Medicago truncatula; Osa, Oryza sativa; Ppa, Physcomitrella patens; Ppe, Prunus persica; Ptr, Populus trichocarpa; Rco, Ricinus communis; Sbi, Sorghum bicolor; Sly, Solanum lycopersicum; Smo, Selaginella moellendorffii; Tur, Triticum urartu; Vvi, Vitis vinifera; Zma, Zea mays. The numbers following the abbreviations are GI numbers. Locus names of Arabidopsis thaliana proteins are shown in parentheses. Asterisks indicate that the protein models were corrected according to the predicted TMD-pSIT architecture in the ORFs (see Materials and Methods for detail). Numbers at the edges of the alignment indicate the first and last (terminal) amino acids of the domains shown. The Phobius-predicted TMD of the first sequence in each alignment is indicated above the sequence. Filled circles indicate proteins predicted not to have a TMD by Phobius. ClustalX color was assigned to the alignments.

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Figure 4.2 Amino acid sequence alignment of the C-terminal domains of predicted plant SIT proteins.

125

Figure 4.3 Amino acid sequence alignment of the C-terminal domains of predicted SIT proteins not verified in this study. (A) – (F) Alignments of predicted plant SIT protein families Full-length proteins were used for alignment (MAFFT with “E-INS-I strategy”) and only the C-termini are shown. Ata, Aegilops tauschii; Bdi, Brachypodium distachyon; Mtr, Medicago truncatula; Osa, Oryza sativa; Ppe, Prunus persica; Ptr, Populus trichocarpa; Rco, Ricinus communis; Sbi, Sorghum bicolor; Tur, Triticum urartu; Zma, Zea mays. (G) Alignment of the predicted animal SIT proteins with known animal KASH proteins. The C-terminal 70 amino acids of each protein were used for alignment (MAFFT with “G-INS-i strategy”). The first two letters in protein names are species name abbreviations: Hs, Homo sapiens; Dm, Drosophila melanogaster; Ce, Caenorhabditis elegans; Ll, Loa loa; Wb, Wuchereria bancrofti; Bm, Brugia malayi ; Bf, Branchiostoma floridae. The numbers following abbreviations are GI numbers. The red lines enclose Phobius-predicted TMDs. Phobius failed to predict the TMD of DmMSP- 300, DmKlarsicht, and CeANC-1. The TMDs of these proteins were determined by the alignment to HsNesprin-1. Red arrows point at newly identified SIT protein candidates. Proteins paired by the red brace are homologs. In (A) – (G), numbers following abbreviations are GI numbers. Numbers at the edges of the alignment indicate the first and last (terminal) amino acids of the domains shown. All other symbols are as in Figure 4.1. ClustalX color was assigned to the alignments.

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Figure 4.3 Amino acid sequence alignment of the C-terminal domains of predicted SIT proteins not verified in this study. 127

Figure 4.4 Subcellular localization of predicted plant SIT proteins. GFP-tagged AtNEMO1, At- NEMO2, AtNEMOL, AtNEMOS, and MsNEME1 under the control of the 35S promoter were stably expressed in wild type (A) or the sun1-KO sun2-KD mutant (C), respectively. GFP-tagged AtNEMO1ΔTVPT, AtNEMO2ΔLVPT, AtNEMOLΔPLPT, AtNEMOSΔLVPT, and MsNEME1ΔLVPT driven by the 35S promoter were stably expressed in wild-type Arabidopsis (B). Root tip cells were imaged using confocal microscopy. Bars equal 5 µm. GFP signal is shown in green. Images in the second column of (A) are overlays of GFP- and transmitted-light images. Cell-to-cell variability of GFP-fusion protein abundance was seen in all images.

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Figure 4.5 Co-IP analysis of AtNEMO-AtSUN interactions. (A) Domain organization of AtNEMO1, AtNEMO2, and their SIT domain mutants. The C4 motif is indicated in bold. (B) Domain organization of AtSUN2 and its mutants. The C-terminal 30 amino acids of AtSUN2 are shown and the residues changed in AtSUN2dMut are indicated in red. Diagrams in (A) and (B) were drawn to scale, with the gaps in (A) representing 300 amino acids. The numbers above each domain indicate the position of the first and the last amino acid of that domain. Blue, domain N-terminal to the TMD; yellow, TMD; white, unknown domain; red, coiled-coil domain; green, N-terminal part of the SUN domain; orange, C-terminal part of the SUN domain. (C) AtNEMO1 and AtNEMO2 interact with AtSUN1 through their SIT domain. (D) AtNEMO1 and AtNEMO2 interact with AtSUN2 through their SIT domain. (E) - (G) AtSUN1 and AtSUN2 interact with AtNEMO1 and AtNEMO2 through their SUN domain. The arrowhead in the bottom- right panel of (E) points at the co-detected GFP-Myc-AtSUN2∆NSUN band. In (C) - (G), GFP-tagged proteins were immunoprecipitated and detected with an anti-GFP antibody. Myc-tagged proteins were detected with an anti-Myc antibody. The input/IP ratio is 1/9. Numbers on the left indicated molecular weight in kD.

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Figure 4.5 Co-IP analysis of AtNEMO-AtSUN interactions.

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Figure 4.6 Co-IP analysis of the AtNEMOL/AtNEMOS/MsNEME1-AtSUN interactions. (A) Domain organization of AtNEMOL, AtNEMOS, MsNEME1 and their SIT domain mutants. Diagrams were drawn to scale. Blue, domain N-terminal to the TMD; yellow, TMD. The numbers above each domain indicate the position of the first and the last amino acid of that domain. The C4 motif is indicated in bold. (B) AtNEMOL interacts with AtSUN1 and AtSUN2 through its SIT domain. (C) AtSUN1 and AtSUN2 interact with AtNEMOL through their SUN domain. (D) AtNEMOS interacts with the SUN domain of AtSUN1 through their SIT domain. (E) AtNEMOS interacts with the SUN domain of AtSUN2 through their SIT domain. (F) MsNEME1 interacts with the SUN domain of AtSUN1 through their SIT domain. (G) MsNEME1 interacts with the SUN domain of AtSUN2 through their SIT domain. In (B) – (G), GFP-tagged proteins were immunoprecipitated and detected with an anti-GFP antibody. Myc-tagged tagged proteins were detected with an anti-Myc antibody. The input/IP ratio is 1/9. Numbers on the left indicated molecular weight in kD.

131

Figure 4.7 F-actin association of AtNEMO11-308 (containing the ARM domain) (A) GFP- AtNEMO11-308 was transiently co-expressed with RFP-Lifeact (1st row) or MAP4-RFP (2nd row) in N. benthamiana leaves, showing that GFP-AtNEMO1 was co-localized with RFP-Lifeact but not MAP4-RFP. GFP-AtNEMO21-309 was transiently co-expressed with RFP-Lifeact in N. benthamiana leaves (3rd row) but they were not co-localized. GFP- AtNEMO11-308 and RFP-Lifeact were co-localized in root cells of stably transformed Arabidopsis plants (4th row). (B) GFP-AtNEMO21-521 was not localized to filament-like structures in transiently expressed N. benthamiana leaves. Scale bars=10 µm.

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Figure 4.8 Expression pattern of AtNEMO1 and AtNEMO2.GUS staining of NEMO1pro::GFP- GUS and NEMO2pro::GFP-GUS transformed wild-type plants (B) Detailed expression pattern of AtNEMO1 and AtNEMO2 and the protein localization in leaves revealed by AtNEMO1pro::GFP- AtNEMO1 and AtNEMO2pro::GFP-AtNEMO2 transformed wild-type plants. For each line, a z- stack image was reconstructed to a 3D image which is shown in different viewing angles. The viewing field is 212.17 µm in height, 212.17 µm in width, and 30 µm in depth. GFP signals are shown in green, and the autofluorescence from chloroplasts is shown in red. AtNEMO1 is expressed in guard cells and guard cell mother cells, while AtNEMO2 is expressed in epidermal cells and mesophyll cells, but very weakly in guard cells. (C) AtNEMO1 is localized to the cytoplasm and enriched at NE fibers in guard cells. (D) The expression of AtNEMO2 is weaker in guard cells than in epidermal cells and mesophyll cells. The images are maximum intensity projections of a z-stack image. Blue arrows indicate a pair of guard cells. (E) Expression pattern of AtNEMO1 and AtNEMO2 and the protein localizations in trichomes, roots, and pollen grains. Both genes are expressed in root cells. AtNEMO1 was enriched at fibers at the root cell NE which was not observed in the localization of AtNEMO2. AtNEMO1 is not expressed in trichomes and pollen grains, while AtNEMO2 is expressed in both cell types and the protein is localized to the NE. In pollen grains, AtNEMO2 is predominantly expressed in sperm cells but weakly expressed in vegetative cells. (F) The AtNEMO1 NE fibers in both root cells and guard cells are sensitive to 1 h treatment of 10 µM LatB. The images of guard cells are maximum intensity projections of a z-stack image. All scale bars equal 10 µm.

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Figure 4.8 Expression pattern of AtNEMO1 and AtNEMO2.

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Figure 4.9 AtNEMO1 is involved in guard cell central nuclear anchorage. (A) T-DNA insertion sites of nemo1-1, nemo1-3, nemo2-1, and nemo2-2. The left border of T-DNA insertion sites were confirmed by sequencing and indicated by arrows on AtNEMO1 and AtNEMO2 genomic structures (drawn to scale). Exons are depicted as thick bars, and introns are depicted as thin lines. DNA fragments encoding the ARM domain and the TMD-SIT domain are shown in red and orange, respectively. (B) Example of measuring nuclear position in guard cells. An ellipse was rendered on a pair of guard cells, and the acute angle between the center of the nucleus and the minor axis (indicated by curved double-headed arrows) was measured. (C) - (E) Distribution curves of nuclear position in different lines. (F) Percentage of the nuclei in the categories divided by the median of wild type (11°, and indicated by a yellow-dotted line). “*”, P<0.05 when compared with wild type; “o”, P>0.05 when compared with wild type; “·”, P<0.01 when compared with nemo1-1 (one-tailed Fisher’s exact test, n=172). The observed numbers are indicated in columns.

135

Figure 4.10 Phenotype comparison between nemo1 and nemo2 mutants and wild type.

136

Primer Sequence Direction Used to clone: 5’-(CACC)ATGGGTGTTGCTGATCTTATTAAGAAGTTCGAGTCTATTT-3’ Forward Lifeact 5’-TCATTCCTCCTTAGAAATAGACTCGAACTTCTTAATAAGA-3’ Reverse Lifeact AtNEMO1, AtNEMO1ΔTVPT 5’-(CACC)(ATG)GGTTTGAATCTGAATCCAATAT-3’ Forward , AtNEMO1XT, AtNEMO11-308 5’-TCATGTAGGGACAGTGTAGTAACCAACA-3’ Reverse AtNEMO1 5’-TCAGTAGTAACCAACATCATCATCT-3’ Reverse AtNEMO1ΔTVPT 5’- TCTGCGGACTTTGTATAAAATCTCACCATCAATATCACAGTTGCGAAG AG-3’ and 5’- Reverse AtNEMO1XT TCAAGCCGCGGCTGCAGCTTCTGCGGACTTTGTATAAAATCTCACCAT CA-3’ 5’-TCAAGTAACAGAACGACAACCTTTCTCC-3’ Reverse AtNEMO11-308 AtNEMO2, AtNEMO2ΔLVPT , AtNEMO2XT, 5’-(CACC)(ATG)GGAAGAAATCTTGGTTCGGCAT-3’ Forward AtNEMO21-309, AtNEMO21-521 5’-TTAAGTCGGAACAAGATGAGGAGGCAT-3’ Reverse AtNEMO2 5’-TTAATGAGGAGGCATCATATCATCC-3’ Reverse AtNEMO2ΔLVPT 5’- TTCTGCGGACTTTGTATAAAATCTGAGATACATCCACATGAAAGAAGC AA-3’ and 5’- Reverse AtNEMO2XT TCAAGCGGCTGCAGCTTCTGCGGACTTTGTATAAAATCTGAGATACAT CC-3’ 5’-TTATCGGGGCTTGCAGTTCTCTGC-3’ Reverse AtNEMO21-309 5’-CAATCACTATTTTAACACACCCTTGCT-3’ Reverse AtNEMO21-521 AtNEMOL, 5’-(CACC)(ATG)AAGGAAATTCAAATTCCCAGG-3’ Forward AtNEMOLΔPLPT , AtNEMOLXT 5’-TCATGTTGGAAGTGGTCCACGGTA-3’ Reverse AtNEMOL 5’-TCATCCACGGTAAGCACGTTCCTCT-3’ Reverse AtNEMOLΔPLPT

continued

Table 4.1 Primers used for cloning. Parentheses enclose the "CACC" for TOPO cloning and the start codon "ATG".

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Table 4.1 continued

5’- CTTCTGCGGACTTTGTATAAAATCTGAAGAAGACCACAATCGCAGCTA AG-3’ and 5’- Reverse AtNEMOLXT TCATGCAGCCGCGGCTGCAGCTTCTGCGGACTTTGTATAAAATCTGAA GA-3’ 5’-(CACC)(ATG)GAAGAAAGAGAGGAGAGTAGTT-3’ Forward AtNEMOS 5’-TCAAGTAGGAACCAAAGTAATGATT-3’ Reverse AtNEMOS AtNEMOSΔLVP 5’-TCAAGTAATGATTTGTGTATTCATGGAG-3’ Reverse T MsNEME1, 5’-(CACC)ACAAATCGTGGAACCCGC-3’ Forward MsNEME1ΔLVP T 5’-GAAAATCAACCTCAGTGAAATGGC-3’ Reverse MsNEME1 MsNEME1ΔLVP 5’-TTATCTACAATCACCATCAGGAGTGAA-3’ Reverse T Flag- 5’-GAAGCGACTCGGCCACTTGCATCTTCCGGTTCAGG-3’ Forward AtSUN1dMut Flag- 5’-CCTGAACCGGAAGATGCAAGTGGCCGAGTCGCTTC 3’ Reverse AtSUN1dMut Myc- 5’-GGAAGCTCTTCAGCCACTTGCATCTTCCGCTTCAG 3’ Forward AtSUN2dMut Myc- 5’-CTGAAGCGGAAGATGCAAGTGGCTGAAGAGCTTCC 3’ Reverse AtSUN2dMut 5’-(CACC)CCAAAAAAGAAGAGAAAGGTAGAAG-3’ and 5’- Forward NLS-GFP-NES AAGAGAAAGGTAGAAGACCCCATGGTGAGCAAGGGCGAG-3’ 5’-TTAATCAAGAGTAAGTCTCTCAAGCGGTG-3’ and 5’- Reverse NLS-GFP-NES TCTCTCAAGCGGTGGTAGCTGAAGCTTGTACAGCTCGTCCATG-3’

138

Primer pair Purpose

5’-CACCATGGGTTTGAATCTGAATCCAATAT-3’ Amplifying the wild-type sequence of AtNEMO1 for nemo1-1 and nemo1-3 5’-TCATGTAGGGACAGTGTAGTAACCAACA-3’ genotyping

5’-TCATGTAGGGACAGTGTAGTAACCAACA-3’ Amplifying the T-DNA sequence of nemo1-1 for nemo1-1 genotyping 5’-TGGTTCACGTAGTGGGCCATCG-3’

5’-TTTGGCTTCTAGTGCTGGTTCTTTC-3’ Amplifying the wild-type sequence of AtNEMO1 for nemo1-2 genotyping 5’-TCATTGAAGAGAGCTGATTAGCTTCC-3’

5’-TTTGGCTTCTAGTGCTGGTTCTTTC-3’ Amplifying the T-DNA sequence of nemo1-2 for nemo1-2 genotyping 5’-TGGTTCACGTAGTGGGCCATCG-3’

5’-CACCATGGGTTTGAATCTGAATCCAATAT-3’ Amplifying the T-DNA sequence of nemo1-3 for nemo1-3 genotyping 5’-ATATTGACCATCATACTCATTGC-3’

5’-CACCATGGGAAGAAATCTTGGTTCGGCAT-3’ Amplifying the wild-type sequence of AtNEMO2 for nemo2-1 and nemo2-2 5’-TTATCGGGGCTTGCAGTTCTCTGC-3’ genotyping

5’-TTATCGGGGCTTGCAGTTCTCTGC-3’ Amplifying the T-DNA sequence of nemo2-1 for nemo2-1 genotyping 5’-TTCATAACCAATCTCGATACAC-3’

5’-CACCATGGGAAGAAATCTTGGTTCGGCAT-3’ Amplifying the T-DNA sequence of nemo2-2 for nemo2-2 genotyping 5’-CCATGTAGATTTCCCGGACATGAAG-3’

Table 4.2 Primers used for genotyping nemo1-1, nemo1-2, nemo1-3, nemo2-1, and nemo2-2.

139

CHAPTER 5

Perspectives

140

Our understanding of plant SUN domain-interacting tail (SIT) proteins and their role in regulating nuclear shape and nuclear positioning has just begun. The research presented in this dissertation revealed the complexity of this novel group of proteins as well as their biological roles and—consequently—leads to a number of further questions about this topic.

5.1 Regulation of nuclear shape and its function

Although the root hair and trichome nuclear shape of wit1-1 wit2-1, wip1-1 wip2-1 wip3-

1, sun1-KO sun2-KD, and kaku1-4 is severely affected, none of these mutants have root hair and trichome developmental defects (Oda and Fukuda, 2011; Tamura et al., 2013;

Zhou et al., 2012a). It has been reported that the elongated root nuclear shape is regulated by the WIT-myosin XI-I complex, which is proposed to be connected to the SUN-WIP complexes. In mammalian cells, mutations of type A Lamins lead to laminopathies (Dahl et al., 2008; Webster et al., 2009). The nuclei of laminopathy patients are often lobulated and invaginated, have altered chromatin organization and an altered nuclear rigidity and sensitivity to mechanical stress. Similar nuclear shape changes occur during aging in C. elegans (Haithcock et al., 2005) and human cells (Scaffidi and Misteli, 2006). In plants, nuclear shape changes have also been observed in the leaf epidermal cells of Arabidopsis nucleoporin nup136 mutants (Tamura et al., 2010) and the root cells of Arabidopsis bearing mutations in long coiled-coil NE protein LINC/CRWN1, 2, and 4 (Dittmer et al.,

2007; Sakamoto and Takagi, 2013). Therefore, it will be important to determine the

141 interaction of these proteins with the SUN-WIP complex and to develop assays that test the effect of mechanical stress on nuclear function in plant epidermal cells.

5.2 Potential motor proteins in pollen nuclear movement

Pollen vegetative nuclear movement is mediated by WIPs and WITs on its NE

(CHAPTER 3). It is known that myosin XI-I is recruited by WITs to the NE and is responsible for the nuclear movement of mature root epidermal cells and mesophyll cells in dark (Tamura et al., 2013). Although myosin XI-I is not responsible for the nuclear movement in pollen tubes (Tamura et al., 2013) (CHAPTER 3), other motor proteins are likely recruited by WITs and regulate the vegetative nucleus movement. Thus, it will be important to understand the protein interactome of WIT1 and WIT2 in pollen.

5.3 Players in nuclear positioning other than SIT proteins

During root hair development, the nucleus is positioned adjacent to the root hair tip.

Similarly, during trichome development, the nucleus migrates to the protruding tip and is usually positioned at the branch points in mature trichomes. None of these processes is interrupted in kaku1-4, wit1-1 wit2-1, witp1-1 wip2-1 wip3-1, wifi, nemo1-1 nemo2-1, nemo1-1 nemo2-2, nemo1-3 nemo2-2, or sun1-KO sun2-KD mutants (CHAPTER 2 and

CHAPTER 4). All these mutants have no whole-plant phenotypes, suggesting that nuclear positioning during asymmetric cell division is not affected. It is possible that these nuclear positioning events are regulated by unidentified plant SIT proteins. It is also possible that these nuclear position events are regulated by other types of proteins.

142

Examples for such proteins in opisthokonts include TOCA-1, which functions to move nuclei in parallel to SUN-KASH bridges in C. elegans P cells (Chang et al., 2012), as well as the RanBP2-BICD2 and Nup133-CENP-F pathways which act sequentially to recruit dynein/dynactin to the nuclear envelope (NE) for the necessary nuclear movement during neuron formation in the mammalian brain (Hu et al., 2013). TOCA-1 and RanBP2 are not conserved in plants, and the N-terminal domain (AA 1-500) of Nup133 which is responsible for recruiting CENP-F (Bolhy et al., 2011) is not conserved in plant Nup133, either. Nonetheless, in plants, it is worth investigating whether nucleoporins or other proteins recruit motor proteins to the NE and regulate nuclear movement.

5.4 Function of AtNEMO1 and AtNEMO2

To my knowledge, nuclear positioning has not been addressed in guard cells. In

CHAPTER 4, I showed that the nuclei are positioned predominantly at the center of guard cells and that AtNEMO1 is required for the full establishment of this positioning.

The biological function of this nuclear positioning is unknown. It is possible that it is related to stomatal aperture or microbe sensing. AtNEMO2 has a different expression pattern from AtNEMO1 in leaves and it is also expressed in sperm cells where AtNEMO1 expression could not be detected. Together with absence of a guard cell nuclear positioning phenotype in nemo2 mutants, this suggests that, despite their sequence similarity, the two proteins have functionally diverged. Further investigation is needed to reveal the detailed functions of AtNEMO1 and AtNEMO2. For example promoter-swap

143 experiments would help dissect the contribution of expression patterns versus protein activities in the divergent roles.

5.5 Potential additional, unidentified SIT proteins

Compared to the nuclear pore proteins, NE proteins appear to be even less conserved between plants and animals. This suggests that the hunt for plant SIT proteins will be predominantly by de novo identification, rather than homology searches. The DORY algorithm provides a good solution to identify possible SIT proteins, but SIT proteins not following the rules implemented in the DORY algorithm may exist. Therefore, protein- interaction screens using the SUN domain are necessary to identify these SIT proteins.

No rapid answers for any of these questions are to be expected, but they will likely lead to a broader, more comparative understanding of the physical interaction of the nucleus with its cellular environment.

144

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