Table of Contents Chapter 1: Introduction

MIAMI UNIVERSITY The Graduate School

Certificate for Approving the Dissertation

We hereby approve the Dissertation

Desheng Liu

Candidate for the Degree: Doctor of Philosophy

______(Dr. Christopher A. Makaroff, Director)

______(Carole Dabney-Smith, Committee Chair)

______(Dr. Michael A. Kennedy, Reader)

______(Dr. David L. Tierney, Reader)

______(Dr. Eileen K. Bridge, Graduate School Representative)

ABSTRACT

OVEREXPRESSION OF NTAP:ATCTF7∆B LEADS TO PLEIOTROPIC DEFECTS IN REPRODUCTION AND VEGETATIVE GROWTH IN ARABIDOPSIS

by Desheng Liu

Eco1/Ctf7 plays a critical role in the establishment of sister chromatid cohesion, which is required for the faithful segregation of replicated chromosomes. Inactivation of Arabidopsis CTF7 (AtCTF7) results in severe reproductive and vegetative growth defects. To further investigate potential roles of AtCTF7 and to identify AtCTF7 interacting proteins, several AtCTF7 constructs were generated and expressed in Arabidopsis plants. 35S:NTAP:AtCTF7∆B (AtCTF7∆299-345) transgenic plants displayed a wide range of phenotypic alterations in reproduction and vegetative growth. Male meiocytes from 35S:NTAP:AtCTF7∆B plants exhibited defective chromosome segregation and ultimately fragmented chromosomes. Mutant ovules developed asynchronously, experienced prolonged meiotic and megagametophytic stages and produced megaspores/embryo sacs that degenerated at various stages. The transgenic plants also exhibited a broad range of vegetative defects, including meristem disruption and apparent epigenetic alterations. Transcripts for epigenetically regulated transposable elements were elevated in transgenic plants. 35S:AtCTF7∆B transgenic plants also exhibited reduced fertility and vegetative defects, with the 35S:AtCTF7∆B defects appearing more severe than those in 35S:NTAP:AtCTF7∆B plants. Additional phenotypes were also observed in 35S:AtCTF7∆B transgenic plants. Therefore, the defects observed in 35S:NTAP:AtCTF7∆B plants are caused by high level expression of AtCTF7∆B and not the presence of the NTAP tag. Finally, Atctf7 plants containing a CTF7pro:AtCTF7∆B construct were obtained and found to grow only slightly better than Atctf7 plants. Therefore, the B motif is required for proper AtCTF7 function. In summary, this study further demonstrates that AtCTF7 plays essential roles in reproduction and in vegetative growth, and that proper levels of AtCTF7 are critical for normal plant growth and development.

OVEREXPRESSION OF NTAP:ATCTF7∆B LEADS TO PLEIOTROPIC DEFECTS IN REPRODUCTION AND VEGETATIVE GROWTH IN ARABIDOPSIS

A DISSERTATION

Submitted to the Faculty of Miami University in partial fulfillment of the requirements for the degree of

Doctor of Philosophy

Department of Chemistry and Biochemistry by Desheng Liu Miami University Oxford, Ohio December, 2014

Dissertation Director: Dr. Christopher A. Makaroff

Table of Contents Chapter 1: Introduction ...... 1 1.1 Mitosis and meiosis...... 1 1.2 Sister chromatid cohesion and chromosome segregation ...... 4 1.3 The cohesin complex ...... 4 1.3.1 The cohesin complex in Arabidopsis ...... 11 1.4 Loading of the cohesion complex on sister chromatids ...... 12 1.4.1 Cohesin loading in Arabidopsis ...... 13 1.5 Cohesion establishment and maintenance ...... 14 1.5.1 Functions of lysine acetylation ...... 16 1.6 Cohesin removal ...... 17 1.6.1 Cohesin removal in mitosis ...... 17 1.6.1.1 Protection and removal of centromeric cohesins ...... 19 1.6.2 Cohesin removal in meiosis ...... 21 1.6.3 Functions of Separase ...... 21 1.7 Cohesion establishment, maintenance and release in Arabidopsis ...... 21 1.7.1 Protection of centromeric cohesion in Arabidopsis ...... 22 1.7.2 Separase in Arabidopsis ...... 23 1.8 Cohesin complex: functions beyond sister chromatid cohesion ...... 24 1.8.1 TranscriptionAL regulation related to the cohesin complex ...... 24 1.8.2 Cornelia de Lange Syndrome (CdLS)...... 25 1.8.3 TranscriptionAL mis-regulation in Roberts Syndrome (RBS) ...... 26 1.8.4 DNA double-strand break repair ...... 27 1.8.5 Human cancers related to mis-function of cohesin ...... 28 1.9 References ...... 30 Chapter 2: 35S:NTAP:AtCTF7∆B leads to various defects in reproduction ...... 48 2.0 Abstract ...... 48 2.1 Introduction ...... 49 2.1.1 Background for Eco1/Ctf7 ...... 49 2.1.2 Esco2 mutations in Roberts Syndrome (RBS) ...... 50 2.1.3 The AtCTF7 protein ...... 50 2.2 Results ...... 56 2.2.1 Experimental design and dominant negative defects of 35S:NTAP:AtCTF7∆B plants ...... 56 2.2.2 35S:NTAP:AtCTF7∆B plants demonstrate defective chromosome segregation during male meiosis ...... 60 2.2.3 35S:NTAP:AtCTF7∆B leads to various defects in ovule development ...... 63 2.2.3.1 Introduction for ovule development in wild type Arabidopsis ...... 63 2.2.3.2 Ovules display various defects due to variations in AtCTF7 expression...... 67 2.2.3.3 35S:NTAP:AtCTF7∆B leads to various defects in ovule development ...... 67 2.2.4 35S:NTAP:AtCTF7∆B leads to disrupted expression in the genes involved in female gametophyte development and other related processes ...... 77 2.3 Discussion ...... 80 2.4 Materials and methods ...... 83 2.5 References ...... 90

Chapter 3: 35S:NTAP:AtCTF7∆B leads to vegetative growth defects ...... 89 3.0 Abstract ...... 96 3.1 Introduction and background ...... 97 3.2 Results ...... 98 3.2.1 Expression of 35S:NTAP:AtCTF7∆B causes pleiotropic growth defects ...... 98 3.2.2 Epigenetic alterations are present in 35S:NTAP:AtCTF7∆B plants ...... 102 3.2.3 Analysis of 35S:AtCTF7∆B and CTF7:AtCTF∆B plants ...... 106 3.3 Discussion ...... 111 3.4 Materials and methods ...... 112 3.5 References ...... 119 Chapter 4: Conclusions and perspectives ...... 121 4.1 References ...... 127

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List of Tables Table 1.1 Cohesin complex subunits and accessory proteins ...... 10 Table 2.1 ECO1/CTF7 from the eukaryotes...... 51 Table 2.2 35S:NTAP:AtCTF7∆B transfer efficiency ...... 58 Table 2.3 Female gametophyte development in WT (Col) plants ...... 69 Table 2.4 Female gametophytes in 35S:NTAP:AtCTF7∆B plants ...... 70 Table 2.5 Primer sequences ...... 87 Table 3.1 Phenotype variations in 35S:NTAP:AtCTF7∆B plants in different generations...... 101 Table 3.2 Primer sequences ...... 115

List of Figures Figure 1.1 Mitosis and meiosis...... 3 Figure 1.2 Architecture of the cohesin complex...... 8 Figure 1.3 The cohesin cycle in eukaryotes (Budding yeast is used as the representive)...... 9 Figure 1.4 Functions of ECO1/CTF7...... 16 Figure 1.5 Cohesion resolution in mitosis...... 20 Figure 2.1ECO1/CTF7 sequences are conserved among eukaryotes...... 52 Figure 2.2 The amino acid sequence of AtCTF7 and the proposed structure of AtCTF7...... 54 Figure 2.3 The 35S:NTAP:AtCTF7∆B construct...... 56 Figure 2.4 35S:NTAP:AtCTF7∆B plants exhibit reduced fertility...... 57 Figure 2.5 Expression analysis of AtCTF7 in wild-type and 35S:NTAP:AtCTF7∆B plants...... 59 Figure 2.6 35S:NTAP:AtCTF7∆B meiocytes demonstrate defective meiotic chromosome organization...... 61 Figure 2.7 Cohesion protein SYN1 distribution pattern is not affected in 35S:NTAP:AtCTF7∆B male meiocytes...... 62 Figure 2.8 Female gametophyte development in WT Arabidopsis plants revealed by differential interference contrast (DIC) microscopy...... 64 Figure 2.9 Female gametophyte development in WT Arabidopsis plants, revealed by confocal laser scanning microscopy (CLSM)...... 66 Figure 2.10 Asynchronous development of early 35S:NTAP:AtCTF7∆B ovules...... 68 Figure 2.11 Female gametophyte development is slowed in 35S:NTAP:AtCTF7∆B plants, revealed by confocal laser scanning microscopy (CLSM)...... 71 Figure 2.12 The meiosis and mitosis of female gametophyte development is slowed in 35S:NTAP:AtCTF7∆B ovules revealed by DIC...... 72 Figure 2.13 35S:NTAP:AtCTF7∆B ovules contain various defective gametophytes during meiosis and mitosis...... 74 Figure 2.14 Female gametophytes from other lines develop slowly and some megaspores alter their identities ...... 76 Figure 2.15 Elevated expression levels of genes involved in female gametophytes in 35S:NTAP:AtCTF7∆B plants...... 78 Figure 2.16 Altered expression levels of genes involved in cohesion, synapsis and recombination in 35S:NTAP:AtCTF7∆B plants...... 79 Figure 3.1 Progressive morphological aggravation of 35S:NTAP:AtCTF7∆B plants through self- pollination ...... 99 Figure 3.2 Phenotypes of 35S:NTAP:AtCTF7∆B dwarf plants...... 100 Figure 3.3 The mRNA levels of the epigenetically regulated transposable elements, MU1, COPIA28 and soloLTR, are dramatically increased in 35S:NTAP:AtCTF7∆B plants...... 103 Figure 3.4 Transcriptional analysis of genes involved in epigenetics, cell cycle and DNA repair...... 104 Figure 3.5 DNA methylation is not completely removed from IgN5 transposons...... 106 Figure 3.6 AtCTF7 constructs used in this study...... 108 Figure 3.7 Morphological variations of 35S:AtCTF7∆B plants...... 109 Figure 3.8 Phenotypes of Atctf7-1-/-plants with CTF7pro:AtCTF7∆B ...... 110

Abbreviations AN: Antipodal nucleus APC/C:Anaphase-promoting complex/Cyclosome At: Arabidopsis thaliana ATP:Adenosine triphosphate ATPase: Adenosine triphosphate nuclease BSA:Bovine serum albumin CARs:cohesin-associated regions CCD: Charged couple device cDNA: Complementary DNA CdLS:Cornelia de Lange Syndrome ChIP:Chromatin immunoprecipitation CLSM: Confocal laser scanning microscopy CN(C): Central nucleus (cell) CTF7: Chromosome transmission fidelity CTCF:CCCTC-binding factor DAPI:4’, 6-diamidino-2-phenylindole DIC: Differential interference contrast DM: Degenerated megaspore DNA:Deoxyribonucleic acid DNAse:Deoxyribonuclease dNTP:Deoxynucleotide triphosphate DSB:Double Strand Break dsDNA:Double stranded DNA ECO1:Establishment of cohesion 1 EN(C): Egg nucleus (cell) FM(L): Functional megaspore (like) GFP:Green fluorescent protein FAA: Formalin, acetic acid and alcohol GUS: β- Glucuronidase HDA:Histone deacetylase

HR:Homologous recombination IP:Immunoprecipitation kDa:Kilodalton mM:millimolar MMC: Megaspore mother cell nm:nanometer mRNA: messenger RNA NCBI: National Center for Biotechnology Information PBS:Phosphate-buffered saline PBST:Phosphate-buffered saline Tween 20 (1XPBS/1mM EDTA/0.1% Tween 20) PCNA:Proliferating cell nuclear antigen PCR:Polymerase chain reaction PFA: Paraformaldehyde Pds5:Precocious dissociation of sisters qRT-PCR: Quantitative reverse transcription polymerase chain reaction Pre-RC:Pre-replication complex rDNA:ribosomal DNA RBS: Roberts Syndrome RFC:Replication factor c RNA:Ribonucleic acid RNAi: RNA interference RT: Reverse transcription SMC3: Structural Maintenance of Chromosome SCC: Sister Chromatid Cohesion SN: Synergid nucleus TAIR: The Arabidopsis Information Resource TEs: Transposable elements T-DNA: Transfer DNA Wapl: Wings apart-like WT: Wild type

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Acknowledgements

First of all, my sincere thanks go to my advisor, Dr. Christopher A. Makaroff, for his continuous encouragement, advice, guidance, patience and support on my research. I would like to thank my committee members, Dr. Carole Dabney-Smith, Dr. Michael A. Kennedy, Dr. David L. Tierney, and Dr. Eileen K. Bridge, who are always kind to offer invaluable suggestion. I would also like to thank all my labmates for their assistance and friendship. Many thanks to Dr. Richard E. Edelmann and Matthew L. Duley for the technical training on microscopy. I am indebted to my parents and my friends for their endless love and support in my life.

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Chapter 1: Introduction

1.1 Mitosis and meiosis

The faithful segregation of the duplicated chromosomes is vital for cell survival during both mitosis and meiosis (Figure 1.1). Mitosis occurs in single-celled prokaryotes during asexual reproduction and in multicellular organisms during growth and development. During mitosis, the chrosmosomes replicate and then separate to produce two genetically identical daughter cells. Mitosis consists of four phases: prophase, metaphase, anaphase and telophase. The DNA replicates during S phase to produce a pair of replicated chromatids. In eukaryotes, the chromosomes are entrapped by sister chromatid cohesion complexes during S stage (Nasmyth, 2001). In prophase, the chromosomes condense and the nucleolus disappears. Then the spindle, which is formed by microtubule fibers, binds the condensed chromosomes and the nuclear membrane breaks down. At metaphase, the chromosomes line up in the middle of the cell for separation. In anaphase, pairs of sister chromatids separate to opposite poles of the cell by the action of the spindle. At telophase, the chromosomes decondense, the nuclear membrane reforms and nucleoli appear. The cytoplasm is separated into two cells by cytokinesis.

Meiosis is responsible for sexual reproduction in eukaryotes. Parent germ cells are duplicated, having pairs of chromosomes. One set of the individual chromosomes are inherited from each parent. During meiosis, the chromosomes duplicate and divide twice to give rise to four gametes, each containing half the number of chromosomes of the original cell. Therefore, meiosis is also called a reduction division. Meiosis consists of two main stages: meiosis I (reductional division) and meiosis II (equational division), consisting of interphase, metaphase, anaphase and telophase. Meiosis II is similar to mitosis, while meiosis I is more complicated as it contains an extra stage, prophase, between interphase and metaphase. During meiosis I the number of chromosomes is reduced to half and genetic material from maternal and paternal chromosomes is exchanged. Prophase I can be further separated into five stages: leptotene, zygotene, pachytene, diplotene and diakinesis (Zickler and Kleckner, 1999; Cnudde and Gerats, 2005; Pawlowski and Cande, 2005). Similar to mitosis, the chromosomes duplicate during S phase before the start of meiosis. During replication, sister chromatid cohesion is established between the chromosomes

via the cohesion complex (Nasmyth, 2001). The chromosomes start to condense at leptotene and homologous chromosomes start to search for each other during late leptotene and early zygotene. The homologous chromosomes start to synapse at zygotene and complete synapsis at pachytene. During thomologous chromosomes synapsis, genetic material is exchanged by crossing over. At diplotene, the homologous chromosomes appear as thick thread-like structures and are tightly associated at chiasmata. At diakinesis, the last stage of prophase I, the chromosomes are further condensed. During metaphase I, the chromosomes are aligned by spindle microtubules on the equatorial plate. At anaphase I, each pair of chromosomes is separated to opposite poles to form two sets of chromosomes, which are still associated at the centromeres. During meiosis II, each daughter cell divides again to give rise to a tetrad of four gametes, which consist of half the number of chromosomes of the original parent cell. Cytokinesis occurs after meiosis II to produce a tetrad of haploid cells. The final germ cells are genetically different than the parent cells as early meiosis results in an increase in genetic diversity. During fertilization, gametes from both parents fuse to produce a diploid set of chromosomes.

Figure 1.1 Mitosis and meiosis. (Left) During mitosis, the duplicated sister chromosomes separate into two genetically identical cells. Mitosis consists of four phases: prophase, metaphase, anaphase, and telophase. (Right) During meiosis, the duplicated chromosomes are separated into four haploid cells. The gametes only contain half the number of chromosomes, which are not identical to the sister chromosomes, of the parent cells. Meiosis consists of meiosis I (reductional division) and meiosis II (equational division). During meiosis I, genetic material is exchanged between the homologous chromosomes.

1.2 Sister chromatid cohesion and chromosome segregation To properly segregate the chromosomes during mitosis and meiosis in eukaryotes, sister chromatid cohesion is established during S phase (Guacci et al., 1994). The duplicated sister chromosomes are maintained by the cohesin complex until they are faithfully segregated into the daughter cells. Sister chromatid cohesion is essential for connection of the kinetochores to the microtubules. At anaphase, cohesion is released and sister chromatids are segregated. In mitosis, cohesion is released in two steps, first from the chromosome arms and then from the centromeres. In meiosis, cohesion is released in two steps. First, cohesion is removed from the homologous chromosome arms to resolve the chiasmata and to segregate homologous chromosomes at diakinesis. Cohesion is then removed from the centromeres to separate the sister chromatids to opposite poles at anaphase II (Lee and Orr-Weaver, 2001; Nasmyth, 2001).

1.3 The cohesin complex The cohesin complex is comprised of six subunits: a heterodimer of Structural Maintenance of Chromosome (SMC) proteins, SMC1 and SMC3, an α-kleisin, either Sister Chromatid Cohesion1 (SCC1/RAD21) in somatic cells or REC8 in meiotic cells, SCC3, Precocious Dissociation of Sisters (PDS5) and WAPL (RAD61) (Figure 1.2). SMC1 and SMC3 are similar in structure and consist of five domains, including an ATP binding domain. The ATPase domain is comprised of the N-terminal and C-terminal domains of each protein. The ATPase domains are connected by α-helical coiled-coil domains, which are linked by a hinge domain (Jones and Sgouros, 2001). SMC1 and SMC3 dimerize to form a heterodimer at the hinge. In some models the ATPase domains of SMC1 and SMC3 also interact with each other (Figure 1.2) (Haering et al., 2004). Mutations in the ATPase domain of SMC1 or SMC3 that cannot hydrolyze ATP cause defects in the ability of the cohesin complex to associate with chromosomes (Arumugam et al., 2003; Arumugam et al., 2006). SCC1/MCD1, the mitotic chromosome determinant, belongs to the α-kleisin (meaning “closure” in Greek) family of proteins (Gruber S. et al., 2003). The N and C termini of SCC1 bind the ATPase domains of SMC3 and SMC1 to form a ring-like structure, which entraps the sister chromatids (Haering et al., 2002; Gruber et al., 2003; Haering et al., 2004). SCC1 helps regulate ATPase activity of SMC1 and SMC3 and is a mediator of the cohesin complex. Due to their essential functions, the N- and C- termini of α-kleisin proteins are

highly conserved (Nasmyth, 2005; Schleiffer, 2003). During meiosis, REC8, a meiosis specific α-kleisin, replaces SCC1 and interacts with SMC1 and SMC3. Genetic studies and biochemical experiments demonstrate that SMC1, SMC3 and SCC1 are the core components of the cohesin complex (Gruber et al., 2003; Losada et al., 1998; Sumara et al., 2000). Disruption of the association of the cohesin complex with chromosomes leads to severe defects. In budding and fission yeast, deletion of Scc1/Mcd1 disrupts chromosome alignment, chromosome condensation and segregation (Guacci et al., 1997; Michaelis et al., 1997; Heo et al., 1998; Tatebayashi et al., 1998). In Drosophila, decreasing Drad21 mRNA by RNAi leads to defects in sister chromatid cohesion and spindle morphology (Vass et al., 2003). In Zebrafish, mutations in Rad21 cause the malfunction of myca, p53 and mdm2 (Rhodes et al., 2010). In budding and fission yeast, deletion of Rec8 disrupts sister chromosome cohesion, homologous chromosome synapsis and axial element formation (Klein et al., 1999). In mouse, inactivating Rec8 causes the loss of sister chromatid cohesion in spermatocytes at meiotic prophase I (Bannister et al., 2004; Tatebayashi et al., 1998). Furthermore, cleavable SCC1 and SMC3 constructs (containing artificial TEV or Separase cleavage sites) have been created to validate their essential roles in cohesion (Gruber et al., 2003; Pauli et al., 2008). SCC3, also known as IRR1 (Irregular) and SA2, is the fourth subunit of the cohesin complex. SCC3 physically interacts with the C-terminus of SCC1 (Haering et al., 2002). In mammalian cells, two Scc3 homologues, SA1 and SA2, have been identified (Losada et al., 2000; Sumara et al., 2000). SA1 and SA2 participate in telomere and centromere cohesion and bind SCC1 exclusively (Canudas and Smith, 2009, Losada et al., 2000; Sumara et al., 2000). SA2 also helps mediate disassociation of the core cohesion complex from the chromosomes during mitosis (Hauf et al., 2005). Mice that are heterozygous for SA1 mutations exhibit tumorgeneisis and increased aneuploidy (Remeseiro et al. 2012). PDS5, WAPL and Sororin also play important roles in cohesin association with the chromosomes. All these proteins play regulatory functions in mediating cohesion maintenance or resolution. Compared to the interactions between the core cohesin subunits, these proteins only transiently associate with cohesin during certain stage(s) of the cohesion cycle. PDS5 is highly conserved among all species (Dorsett et al., 2005; Hartman et al., 2000; Losada et al., 2005; Panizza et al., 2000; Stead et al., 2003; Sumara et al., 2000; Tanaka et al., 2001; Wang et al., 2002). PDS5 interacts with SCC1 and participates in cohesin removal during

early mitosis (Shintomi and Hirano, 2009). PDS5 interacts sub-stoichiometrically with the core subunits of the cohesin complex (van Heemst et al., 1999) and interacts with chromatin through interaction with SCC1 (Mc Intyre et al., 2007). In budding yeast, C. elegans and Drosophila, Pds5 is required for sister chromatid cohesion maintenance and chromosome condensation (Hartman et al., 2000; Panizza et al., 2000; Dorsett et al., 2005). Higher eukaryotes contain more than one Pds5 protein (Losada et al., 2005; Sumara et al., 2000). In vertebrates, there are two PDS5 isoforms, PDS5A and PDS5B. In Xenopus and human cells, deletion of one isoform of Pds5 does not lead to severe defects, even though it results in partial cohesion loss and minor defects in mitotic chromosome assembly. Deletion of both Pds5A and Pds5B in Xenopus cells leads to reduced cohesin levels at centromeres, but increased cohesin levels on the chromosome arms (van Heemst et al., 1999). WAPL (wings apart like) was first discovered in Drosophila and shown to be important for larval development and wing formation (Verni et al., 2000). WAPL binds chromatin from telophase to prophase and interacts with the cohesin complex during all stages of cohesion cycle. The N-terminus of WAPL is variable and less conserved, while the C-terminus is well conserved and forms a helical repeat domain. In animals, the N-terminus contains three FGF motifs, which interact with PDS5 and SCC1. However, FGF motifs are not present in the yeast or plant proteins (Shintomi and Hirano, 2009). In some models, WAPL binds both PDS5 and SCC3 to form a complex to prevent the establishment of cohesion (Rowland et al., 2009). WAPL is also evolutionarily conserved. It is involved in cohesin dissociation from the chromosome by physically interacting with the cohesin complex (Gandhi et al., 2006; Kueng et al., 2006, Ben- Shahar et al., 2008; Rowland et al., 2009; Shintomi and Hirano, 2009;Sutani et al., 2009; Bernard et al., 2008). In addition to its essential roles in cohesin removal, WAPL also promotes chromosome segregation and heterochromatin formation (Dobie et al., 2001; Perrimon et al., 1985; Verni et al., 2000). In budding yeast, Wapl mutations lead to cohesion defects in heterochomatic regions (Verni et al., 2000). In Drosophila, Wapl inactivation leads to the formation of unusual X-shaped metaphase chromosomes and chromosome cohesion loss in heterochomatic regions (Warren et al., 2004). In HeLa cells, Wapl inactivation does not affect chromosome segregation, but increases the resident time of cohesin on chromosomes (Kueng et al., 2006). Meanwhile, cell cycle progression is checked at G2/M and nuclear morphology is altered in the HeLa cells (Gandhi et al., 2006; Kueng et al., 2006).

Sororin has only been identified in vertebrates, where it is essential for sister chromatid cohesion (Rankin et al., 2005; Schmitz et al., 2007). Sororin is a cell-cycle regulated protein, which is degraded by the Anaphase-promoting complex/Cyclosome (APC/C) during the Growth 1/Gap 1 phase (G1 phase). During cohesion establishment, Sororin replaces WAPL to form a stable complex with PDS5. After it is phosphorylated at prophase, Sororin comes off the chromosomes to help remove cohesin from the chromosome arms. The cohesion process is comprised of several steps, including cohesin loading to the chromatin, cohesion establishment, cohesion maintenance and cohesion resolution (Figure 1.3). Every step of the process is regulated by proteins and the cohesin complex is finely tuned by modifications, such as acetylation. Not surprisingly, the general cohesion mechanism and cohesin proteins are conserved in eukaryates (Table 1.1) (Yuan et al., 2011).

Figure 1.2 Architecture of the cohesin complex. The ring-like structure of the cohesin complex embraces the chromosomes. The cohesin complex is composed of six subunits: SMC1, SMC3, an α-kleisin, SCC3, PDS5 and WAPL. SMC1 and SMC3 contain ATPase domains at the termini. The ATPase domain and the hinge domain of SMC1 or SMC3 are linked by α-helical coiled-coil domains. The ATPase domains of SMC3 and SMC1 bind the N- and C termini of an α-kleisin respectively. The α-kleisin directly or indirectly associates with SCC3, PDS5 and WAPL.

Figure 1.3 The cohesin cycle in eukaryotes (Budding yeast is used as the representive). The cohesion cycle is comprised of cohesin loading, cohesion establishment, cohesion maintenance and cohesion resolution. The cohesin complex is loaded at late G1 with the help of SCC2/SCC4, the loading complex. Cohesion is established by ECO1/CTF7 during S phase as ECO1/CTF7 acetylates the ATPase domain of SMC3 and other factors. Cohesin is removed at anaphase when α-kleisin is cleaved by Separase and the sister chromatids are segregated to opposite poles of the cells. The cohesin complex is removed from the chromosomes and deacetylated for recycling (Uhlmann, 2004).

Table 1.1 Cohesin complex subunits and accessory proteins (Yuan et al., 2011).

S.cerevisiae S.pombe C.elegans D.melan Vertebrates A. thaliana Rice Maize

ogaster

Cohesin subunits SMC1 PSM1 HIM-1 SMC1 SMC1A SMC1 SMC1

SMC1B

SMC3 PSM3 SMC-3 SMC3 SMC3 SMC3 SMC3

SCC1/ RAD21 SCC- RAD21 RAD21 SYN2/RAD21.1 RAD21-1 1/COH-2 MCD1 SYN4/RAD21.3 RAD21-2 COH-1 SYN3/RAD21.2 RAD21-3 COH-3 RAD21-4

REC8 REC8 REC8 C(2)M REC8 SYN1/DIF1 AFD1

SCC3/ REC11 SCC3 SA1 SA1/SA2 SCC3 SCC3

IRR1 STAG3

Loading SCC2 MIS4 NIPBL NIPPED-B NIPBL SCC2 OSJ_22834

OSI_24633

SCC4 SST3 MAU-2 MAU-2 SCC4/MAU2 SCC4

Establish ECO1/ ESO1 DECO ESCO1 CTF7 OSI_19739 ESCO1 ment CTF7 ESCO2

Maintenace PDS5 PDS5 ELV-14 PDS5 PDS5A At1g15940

PDS5B At1g77600

At1g80810

At4g31880

At5g47690

Rad61/ WPL1 WAPL-1 WAPL WAPAL At1g11060 OsI_34181 Acl54412

wpl1 At1g61030

Dissociation PDS1 CUT2 IFY-1 PIM SECURIN/PTTG

ESP1 CUT1 SEP-1 SSE SEPARASE/ESP AtESP1 Osi_09098 L1 OsI_08533

CDS5 PLO1 POLO POLO PLK1

SGO1 SGO2 SGO1 MEI-S332 SGO2 AtSGO1 OsJ_08740 ZMSGO2

SGO1 SGO1 AtSGO2 OsI_09305 ZMSGO1

PANS1

PANS2

1.3.1 The cohesin complex in Arabidopsis SMC1 and SMC3 are identified as single-copy genes in the Arabidopsis genome. Arabidopsis thaliana SMC1 (AtSMC1) and AtSMC3 are conserved and share the same functional structures as other SMC proteins: an N-terminal ATP binding domain and a C-terminal DA box linked by two antiparallel α-helical coiled coil regions. AtSMC3 is found in the cytoplasm and nucleus of somatic and germ cells. AtSMC3 localizes on sister chromatids from prophase to anaphase and on the spindles from metaphase to telophase (Lam et al., 2005). Until now, AtSMC1 immunolocalization has not yet been carried out. Inactivation of AtSMC3 (titan 7-1 and titan 7-2) and AtSMC1 (titan8-1 and titan 8-2) by T-DNA insertion leads to embryo and endosperm defects, and the arrest of seeds at early developmental stages (Liu et al, 2002; Lam et al., 2005). Plants heterozygous for mutations in SMC1 or SMC3 display a reduction in the positional alignment of sister chromatids, even though plant morphology is normal and full length transcript levels are approxmiately 77% of wild type plants (Schubert et al, 2009). Four α-kleisins, SCC1/REC8 orthologs exist in the Arabidopsis genome: SYN1, SYN2, SYN3 and SYN4. SYN1/DIF1 only functions in meiosis and is the REC8 ortholog (Bai et al, 1999; Peirson et al, 1997; Cai et al, 2003; Bhatt et al, 1999). Like other REC8 orthologs, SYN1 is essential for sister chromatid cohesion and pairing of homologous chromosomes during meiosis. In wild type plants, SYN1 specifically localizes on the meiotic chromosome axes from leptotene to metaphase I (Cai et al., 2003). SYN1 dissociates from the chromosome arms during diplotene and diakinesis, and specifically localizes at the centromeres at early metaphase I. SYN1 is not detected on the chromosomes by late metaphase I. Homozygous syn1 plants are sterile (Cai et al., 2003). The male meiocytes display severe defects in sister chromatid cohesion, homologous chromosome pairing and chromosome condensation (Bai et al, 1999; Peirson et al, 1997; Cai et al, 2003; Bhatt et al, 1999). Unexpectedly, recombination appears to be partially normal in syn1 mutants (Zhao et al, 2006). Vegetative growth is also normal in syn1 plants. The other three α-kleisins, SYN2, SYN3 and SYN4, are expressed in all plant tissues with the highest levels in meristematic regions (Dong et al., 2001). Among them, SYN2 and SYN4 are the SCC1 orthologs and specifically function in mitosis. Immunolocalization studies show SYN2 and SYN4 label the chromosomes from interphase to late mitotic prophase. SYN2 also functions in DNA repair (da Costa-Nunes et al., 2006). Inactivation of SYN2 or SYN4 does not lead to any

apparent defect in vegetative growth, flowering or seed setting. Syn2-1syn4-1 double homozygous plants are not obtained and about 50% of syn2-1syn4-1+/- seeds are embryo lethal indicating SYN2 and SYN4 have overlapping functions in mitosis (Liu, 2005). SYN3 is an atypical α-kleisin. It primarily localizes in the nucleolus of both somatic cells and male meiocytes and is essential for megagametogenesis (Jiang et al., 2007). The nucleolar localization of SYN3 suggests that SYN3 is not a component of the typical cohesin complex and may function in rDNA structure, rRNA processing and transcription. Homozygous T-DNA syn3-/- plants are not detected. Heterozygous Syn3+/- plants are normal in vegetative growth but produce shorter siliques with fewer ovules. Atsyn3-2- embryo sacs develop asynchronously and contain various defects, including variable numbers of nuclei at FG6, nuclear migration defects and delayed senescence. Defects are also observed after the ovules are fertilized. SYN3 is also essential for meiosis and altered transcription patterns in genes involved in synapsis and recombination are observed if SYN3 expression is disturbed (Yuan et al., 2012). The Arabidopsis genome also contains a single copy of AtSCC3, which is universely expressed in all tissues (Chelysheva et al, 2005). AtSCC3 localizes on the chromosomes during all stages of mitosis and on the chromosome axes until anaphase I in meiosis (Storlazzi et al, 2003). Homozygous Atscc3-1 plants are severely dwarf and sterile. A truncated AtSCC3 protein is detected in Atscc3-1 plants. The meristem regions of Atscc3-1 plants contain fewer numbers of dividing cells. Homozygous Atscc3-1 plants produce aborted seeds with severe embryo defects (Chelysheva et al, 2005). In Atscc3-1 male meiocytes, chromosome alignment and chromosome condensation are disrupted. Unexpectedly, SYN1 protein labeling is normal in Atscc3-1 male meiocytes. Similar to the situation for plants heterozygous for mutations in SMC1 or SMC3, plants heterozygous for SYN3, SCC2 or SCC3 also display a reduction in the positional alignment of sister chromatids while plant morphology is normal and full length protein levels are not dramatically reduced (Schubert et al, 2009).

1.4 Loading of the cohesion complex on sister chromatids The cohesin complex is loaded on the chromosomes by the loading complex, SCC2/SCC4, at the G1 stage (Figure 1.3). Without the help of SCC2/SCC4, cohesin cannot bind the centromeres or chromosome arms (Nasmyth and Haering, 2009; Onn et al., 2008; Ciosk et al., 2000; Gillespie

et al., 2004; Watrin et al., 2006). After the cohesin complex is recruited to the right locus, the hinge domains of SMC1 and SMC3 temporarily open to entrap the chromosome (Gruber et al., 2006). Alternatively, SCC2/SCC4 may modify the chromatin to facilitate cohesin binding (Hakimi et al., 2002; Huang et al., 2004; Ritchie et al., 2008). The original loci where the cohesin complexes are recruited to the chromosome may not be their final destinations. Cohesin complexes may first bind at specific loci on chromosomes (Kogut et al., 2009; Misulovin et al. 2008). Then the loaded cohesins are moved to other loci by RNA polymerases (Lengronne et al., 2004; Hu et al., 2011). As mapped in seveal organisms, the binding loci of cohesin and SCC2 are non-random and not totally overlapping (Blat et al., 1999; Bernard et al. 2001; Nonaka et al., 2002; Glynn et al., 2004; Weber et al., 2004; Lengronne et al., 2004; Schmidt et al., 2009; D’Ambrosio et al., 2008). After loading the cohesin complex, SCC2/SCC4 activates the SMC ATPase activity to modify the cohesin complex and facilitate interaction between the cohesin complex and the chromosomes (Haering et al., 2002; Arumugam et al. 2003; Gruber et al., 2003; Gruber et al., 2006; Seitan et al. 2006). Not surprisingly, mutations in the ATPase domains of SMC1 or SMC3 result in phenotypes that resemble SCC2/SCC4 mutants: the association of cohesin with chromosomes is disrupted (Arumugam et al., 2003). Other factors such as the Cdc7/Drf1 kinase (DDK), kinetochore proteins and tRNA transcription factors are also required for cohesin binding. DDK is a component of pre-replication complexes (pre-RCs), which help mediate cohesin loading (Ström et al., 2007; Takahashi et al., 2004; Gillespie and Hirano, 2004). The kinetochore and tRNA transcription factors also facilate cohesin loading (Weber et al. 2004; D’Ambrosio et al. 2008). Even though the SCC2/SCC4 complex is required for cohesin loading, it is not required for cohesin maintenance and resolution (Ciosk et al., 2000; Lengronne et al., 2006).

1.4.1 Cohesin loading in Arabidopsis Single copies of Scc2/Scc4 are found in the Arabidopsis genome (Sebastian et al. 2009). In Arabidopsis, AtSCC2 plays essential roles in sister chromatid cohesion establishment. Decreasing AtSCC2 mRNA levels by RNAi results in chromosome dis-organization and alterations in the binding of AtSCC3 during meiosis. T-DNA mutations in AtSCC2 or AtSCC4 lead to defective embryo and endosperm development (Sebastian et al., 2009). Compared to non- plant SCC2 proteins, AtSCC2 contains huntingtin-elongation-A subunit-TOR (HEAT) repeats

and an extra plant homeodomain (PHD) finger, which is predicted to function in gene regulation and chromatin organization. Studies on the interaction between AtSCC2/AtSCC4 and the cohesin complex and localization studies of AtSCC2/AtSCC4 have not been carried out.

1.5 Cohesion establishment and maintenance Sister chromatid cohesion is established by ECO1/CTF7 during S phase (Figure 1.3), even though the cohesin complex binds the chromatin in interphase (Uhlmann and Nasmyth, 1998). Cohesion establishment occurs when ECO1/CTF7 acetylates conserved lysine residues in SCC1 and SMC3, which are close to the ATPase domain (Figure 1.4) (Ben-Shahar et al., 2008; Rowland et al., 2009). Acetylated SMC3 stabilizes the interaction between the cohesin complex and the chromosome, and counteracts the disassociation activity of RAD61/WAPL. In animal cells, acetylated SMC3 recruits Sororin, which replaces RAD61/WAPL from the PDS5-cohesin complex to further stabilize the cohesin-chromosome complex (Nishiyama et al., 2010; Schmitz et al., 2007). Now the sister chromatid cohesion complex is competent and the cohesin/chromosome complex is stable. Mutations in ECO1CTF7 lead to various phenotypes including sister chromatid cohesion defects, the altered distribution of cohesin proteins, cell cycle check point activation and growth retardation. During the cohesion establishment process, ECO1/CTF7 interacts with various factors. ECO1/CTF7 physically interacts with the three RFCs (components of the clamp loader replication factor C), which play different roles during replication (Kenna and Skibbens, 2003). ECO1/CTF7 also associates with Ctf4 and Ctf18, which interact with the replication fork and are essential for cohesion (Lengronne et al., 2004). ECO1/CTF7 physically interacts with PCNA (DNA polymerase processivity factor) in budding yeast and human (Moldovan et al., 2006). The sumoylation of PCNA actually suppresses cohesion (Moldovan et al., 2006). DNA helicase also interacts with ECO1/CTF7 (Skibbens, 2004). Sister chromatid cohesion can also be established during G2 phase when double-strand breaks (DSBs) occur. Sister chromatid cohesion is required for DSB repair and the post-replicative DNA repair pathway is defective in eco1 mutants (Strom et al., 2007; Unal et al., 2004; Unal et al., 2008). After DSBs occur, the histone H2AX is phosphorylated, which recruits cohesin complexes to DSB lesions and CARs (Unal et al., 2004). DNA damage also stimulates Chk1 kinase activity to phosphorylate cohesin’s α-kleisin subunit and stimulate ECO1/CTF7 to acetylate the cohesin subunits (Heidinger-Pauli et al., 2008;

Heidinger-Pauli et al, 2009; Kim et al., 2010). The DSB triggered cohesion establishment during G2 phase requires the acetyltransferase activity of ECO1/CTF7. Cohesion can also occur when ECO1/CTF7 is overexpressed during G2 phase without DSBs (Unal et al., 2007). Unlike the cohesion establishment in S phase, cohesion establishment at G2 phase does not require DNA replication. During this process, sister chromatid cohesion is established both on the damaged DNA and on normal DNA. The sister chromatid cohesion mechanism during S phase is different compared to G2 phase. DSBs activate the checkpoint kinase Mec1/ATR, which then triggers the Chk1 kinase. Activated Chk1 then phosphorylates SCC1 at Ser83. The phosphorylated Ser83 in SCC1 mediates the ECO1 mediated acetylation (Unal et al., 2007), during which Lys84 and Lys210 of SCC1 are acetylated. Cells that contain a nonphosphorylatable S83A mutant of SCC1 or nonacetylatable arginines (K84R, K210R) display similar phenotypes; the cohesin complex can bind the chromosomes around the DSBs and at CARs, but cohesion cannot be established. Alternatively, the acetyl-mimic glutamines (K84Q, K210Q) of SCC1 induce cohesion without the involvement of ECO1. In addition to phosphorylation and acetylation, SCC1 also undergoes sumoylation when DSBs occur (McAleenan et al., 2012; Almedawar et al., 2012), which is also essential for DSBs-triggered cohesion.

Figure 1.4 Functions of ECO1/CTF7. Sister chromosome cohesion is established by ECO1/CTF7, which acetylates conserved lysine residues in SCC1 and SMC3, close to the ATPase domain. The acetylated SMC3 causes a conformational change that blocks the dissociation activity of RAD61/WAPL to form a stable complex between the chromosome and cohesin.

1.5.1 Functions of lysine acetylation During cohesion establishment, ECO1/CTF7 acetylates conserved lysine residues of SMC3 to stabilize the interaction between the cohesin complex and chromosomes (RolefShahar et al., 2008; Unal et al., 2008; Zhang et al., 2011). In budding yeast, immunoprecipitation (IP) experiments show that K112 and K113 are acetylated by ECO1 in vivo (Rowland et al., 2009). These lysine residues are essential for yeast survival, as point mutations in these residues make yeast inviable (Rowland et al., 2009). In other eukaryotes, the lysine residues corresponding to yeast K112 and K113 are conserved and acetylated during cohesion establishment (Zhang et al., 2008a). The SMC3 acetylation state changes during the cell cycle with the highest level occuring in S phase and the lowest level at anaphase (Skibbens, 2009; Beckouet et al., 2010; Rowland et al., 2009). SMC3 acetylation by ECO1 is important for cohesion establishment during S phase as

acetylated SMC3 stablizes the cohesin/chromosome interaction and opposes the destabilising interaction of WAPL/PDS5. SMC3 acetylation also depends on the SCC2/SCC4 complex and the replication forks, especially the prereplication protein cell division control protein 6 (Cdc6; Ben- Shahar et al., 2008). SMC3 is deacetylaed by the histone deacetylase Hos1 during anaphase when SCC1 is cleaved and cohesin dissociates from chromatin (Figure 1.3) (Borges et al., 2010). Cells containing a Hos1deletion or SMC3 K112N/K113N mutation display severe cohesion defects. Therefore, the deacetylation of SMC3 is also important, because only deacetylated SMC3 is used as the substrate for the next cycle of cohesion establishment. Interestingly, deletion of Rad61/Wapl or mutations in specific domains of PDS5, SMC3 or SCC3 suppresses the lethal nature of ECO1/CTF7 mutations (Rowland et al., 2009). In addition, deletion of Wapl also rescues the non-acetylatable K112R/K113R mutations of Smc3 (Rowland et al., 2009). The rescue experiments suggest that acetylation activity of ECO1 can overcome the destabilizing effects of WAPL, PDS5 or SCC3 (Rowland et al., 2009). Acetylation of cohesion factors by ECO1/CTF7 may disrupt their functions in cohesion establishment during S phase. An ‘antiestablishment’ complex might be formed by WAPL, PDS5 and SCC3 to maintain the cohesin complex in an unstable state as a WAPL, PDS5 and SCC3 complex has been isolated from budding yeast extractS (Rowland et al., 2009). In this way, ECO1/CTF7 cohesion establishment activity is dispensable without the anti-establishment functions of other proteins. Alternatively, the modified cohesin factors may counteract ECO1/CTF7 cohesion establishment activity (Sutani et al., 2009; Ben-Shahar et al., 2008; Heidinger-Pauli et al., 2008; Rowland et al., 2009; Unal et al., 2008; Zhang et al., 2008a; Shintomi and Hirano, 2009). Meanwhile, the loss of SMC3 acetylation in postreplicative cells does not affect the association of cohesin with chromatin.

1.6 Cohesin removal 1.6.1 Cohesin removal in mitosis In order to separate the duplicated chromosomes into daughter cells, the cohesins need to be removed from the chromosome arms and the centromeres. Cohesin removal requires various factors, such as WAPL, PDS5, Sororin, SGO1 and PAN1. Cohesin removal in mitosis and meiosis is different and some different factors are involved. In mitosis, cohesin removal occurs in two steps, during prophase for chromosome condensation facilitated by WAPL and at

metaphase/anaphase assisted by Separase. In mitosis, cohesin is sequentially removed first at prophase and then at anaphase (Figure 1.5). During prophase, cohesin complexes are removed without being cleaved or degraded, and the undamaged cohesin complexes may be reused for the next cell cycle (Wendt et al., 2008). During prophase, Polo-like kinase 1 (Plk1), which is recruited to cohesin by Sororin at the chromosome arms, and Aurora B phosphorylate SCC3 (Losada et al., 2002; Sumara et al., 2002; Gimenez- Abian et al., 2004). SCC1 protein on the chromosome arms is also phosphorylated and then cohesin is removed from the chromosomes in a Separase-independent process (Waizenegger et al., 2000; Losada et al., 2002). During prophase, cohesins at the centromeres and at the interface between sister chromatid axes are protected and resistant to removal (Waizenegger et al., 2000; Losada et al., 2000). In Xenopus cells, decreasing Plk1 leads to delayed dissociation of cohesin from the chromosomes (Sumara et al., 2002). Expression of a phosphorylated mutant of SA2 promotes the dissociation of cohesin from the chromosomes as early as interphase, even when Plk1 is deleted. Expression of a non-phosphorylatable form of SA2 reduces cohesin dissociation from chromosomes (Zhang et al., 2011). The phosphorylation of SA2 is more crucial than the phosphorylation of SCC1. Expression of a non-phosphorylatable form of SCC1 does not significantly reduce cohesin dissociation (Zhang et al., 2011). Sororin is phosphorylated by Cdk1 and the phosphorylation is essential for the interaction with Plk1. Expressing a nonphosphorylatable form of Sororin (T159A) leads to defects in cohesion resolution at chromosome arms (Losada et al., 2002). Aurora B also regulates condensin I and Sgo1 for chromosome binding (Lipp et al., 2007). In addition to the phosphorylation functions of Plk1, WAPL is also required for cohesin removal during prophase. After cohesin removal in animal cells, WAPL replaces phosphorylated Sororin to form a Pds5-Wapl complex, which destabilizes putative cohesin-chromosome interactions (Schmitz et al., 2007; Nishiyama et al., 2010). Inactivation of Wapl leads to prolonged residence of cohesin on chromosomes and severe defects in cohesion resolution at the chromosome arms, which is more severe than the knockout of Plk1, Aurora B or Condensin I (Kueng et al., 2006). In contrast, Wapl overexpression promotes the separation of sister chromatids (Gandhi et al., 2006). The functions of WAPL in cohesin removal is related to the phosphorylation functions of Plk1, Aurora B and condensin I, as Wapl inactivation does not alter the phosphorylation state of SA2 or affect the localization patterns of other related proteins

(Holm and Rosenstrom, 2010). During prophase, the chromosome arms may be untangled to help the condensation of chromosomes, which promotes topoisomerase meidated decatenation (Holm and Rosenstrom, 2010).

1.6.1.1 Protection and removal of centromeric cohesins Before anaphase, cohesin complexes on the centromeres are protected from phosphorylation by Shugoshin (Sgo) protein (Watanabe, 2005; Salic et al., 2004; Kitajima et al., 2005; McGuinness et al., 2005). Other factors, including PP2A (phosphatase 2A), BUB1, Aurora B, Haspin (a histone H3 kinase) and PHB2 (Prohibitin 2), are also involved in cohesin protection at the centromeres (Dai et al., 2006). SGO1 is recruited to the centromeres by BUB1, the spindle checkpoint protein (Tang et al., 2004; Kitajima et al., 2005), and by Heterochromatin protein HP1 (Yamagishi et al., 2008). Depletion of Budding Uninhibited by Benzimidazoles 1 (Bub1) or heterochromatin protein 1 (Hp1) leads to Shugoshin1 (SGO1) removal from the centromeres and sister chromatid cohesion loss. SGO1 protects the centromeres from the beginning of mitosis. In mammalian cells, inactivation of SGO1 promotes cohesin dissociation and sister chromatid separation before anaphase (Kitajima et al., 2004). In these cells, cohesins at centromeres are removed without the involvement of activated APC/C and Separase. SGO1 physically interacts with PP2A to recruit it to the centromeres for SA2 dephosphorylation in mammalian cells (Tang et al., 2006). The recruited PP2A at centromeres protects cohesin from Plk1-mediated phosphorylation. Expressing a nonphosphorylatable SA2 rescues the Sgo1 deletion mutant (McGuinness et al., 2005). At metaphase, the chromosomes are correctly bi-oriented and cohesins at centromeres are removed at anaphase. At anaphase, the anaphase-promoting complex (APC/C) is activated to cleave the ubiquitylated Securin and Cyclin B (Michaelis et al., 1997; Hauf et al. 2001; Musacchio and Salmon, 2007; Uhlmann et al., 1999). Cleavage of Securin releases Separase which becomes activated (Hornig et al., 2002; Waizenegger et al., 2002). Meanwhile, SGO1 is phosphorylated by Aurora B/Ipl1 and removed from the centromeres, and SCC1 at the centromeres is phosphated by Plk1 (Alexandru et al., 2001). SCC1 is then cleaved by the activated Separase and cohesins at the centromeres are removed (Uhlmann et al., 1999; Waizenegger et al., 2000; Losada et al., 2002; Sumara et al., 2000). The cleaved SCC1 fragments are ubiquitylated and degraded (Rao et al., 2001). Released SMC3 is reused after deacetylation

by histone lysine deacetylase 1 (Rivera and Losada, 2009). Mutations in Separase lead to chromosome segregation defects (Hauf et al., 2001).

Figure 1.5 Cohesion removal in mitosis. In mitosis, cohesin is removed at prophase and at anaphase. During prophase, SCC1 on the chromosome arms is phosphorylated and cohesin complexes at the chromosome arms are removed in a Separase-independent process. Meanwhile, cohesins at the centromeres are protected by Sgo1-PPA2. At metaphase, cohesins at the centromeres are phosphorylated, which disrupts their interaction with SGO1, allowing them to be cleaved by Separase.

1.6.2 Cohesin removal in meiosis Cohesin removal in meiosis also occurs in multiples stages. During diakinesis, cohesins at the chromosome arms are removed, but cohesins at the centromeres remain intact until metaphase II. During anaphase II, the sister chromatids separate as the cohesion at centromeres is cleaved by separase. Compared to mitosis, the homologous chromosomes undergo crossing over in meiosis. The homologous chromosomes are resolved with the disassembly of the synaptonemal complex (SC) in diplotene and the release of cohesins from the chromosome arms at anaphase I. Similar to mitosis, cohesins at pericentromeric regions are protected by SGO1 with the association of PP2A. Dephosphorylated REC8 at centromeric regions is also protected by PP2A and SGO1 (Riedel et al., 2006). Centromeric cohesion is also protected during anaphase I and in the interkinesis (Prieto et al., 2001; Revenkova et al., 2001). During metaphase II, SGO-PP2A complexes disassociate from the centromeres and REC8 is cleaved by Separase. At the end of meiosis II, each daughter cell receives a complete haploid genome from the diploid mother cell.

1.6.3 Functions of Separase The cleavage of SCC1/REC8 requires an activated Separase. Mutations in SCC1/REC8 cleavage sites prevent cohesin removal from the chromosomes (Uhlmann et al., 1999; Uhlmann et al., 2000). Before anaphase, Separase is inhibited by Securin (PDS1 in S. cereviae; Cut2 in S. pombe) and Cyclin B (Yanagida, 2000; Gorr et al., 2005). At the beginning of anaphase, Securin and Cyclin B are ubiquitined and degraded by the anaphase-promoting complex/cyclosome (APC/C) (Irniger et al., 1995; Cohen-Fix et al., 1996; Funabiki et al., 1996; Yamamoto et al., 1996; Uhlmann, 2003; Nasmyth and Haering, 2005). Separase is then released from the Separase-Securin complex and becomes active. In mitosis, activated Separase cleaves SCC1, allowing the release of sister chromatid cohesion (Ciosk et al., 1998; Uhlmann et al., 1999). In meiosis, the activated Separase cleaves REC8 along the chromosome arms during meiosis I and phosphated REC8 at centromeres at anaphase II (Buonomo et al., 2000).

1.7 Cohesion establishment, maintenance and release in Arabidopsis The establishment and maintenance of sister chromatid cohesion are conserved in Arabidopsis. AtCTF7 is able to complement the temperature-sensitive yeast ctf7-203 mutant and is essential

for sister chromatid cohesion establishment (Jiang et al., 2010). The functions of AtCTF7 are fully addressed in Chapter 2. In non-plant organisms, the effects of Wapl inactivation are observed in mitosis but nothing was known about its roles in meiosis. Recently, the function of Arabidopsis WAPL has been characterized. There are two WAPL genes in Arabidopsis and they function in cohesin removal during meiotic prophase (De et al., 2014). While homozygous mutants for either WAPL gene do not show any obvious phenotype, double homozygous wapl plants display reduced fertility and severe defects in male meiosis, including heterochromatin dissociation defects, chromosome bridges and uneven chromosome segregation. In contrast, cohesin removal in mitosis is relatively normal (De et al., 2014). Just like the situation in other species, inactivation of the two Arabidopsis WAPL genes rescues AtCTF7 inactivation. Homozygous Atctf7 plants are severely dwarf and sterile. Wapl1wapl2Atctf7 triple homozygous plants are normal in vegetative growth and produce some viable seeds (De, Bolaños-Villegas and Makaroff, unpublished). SYN1 immunolocalization studies demonstrate that cohesin removal from the chromosomes is recovered in wapl1wapl2Atctf7 triple homozygous mutant plants (De, Bolaños-Villegas and Makaroff, unpublished). Five putative PDS5 genes have been identified in the Arabidopsis genome. Plants single homozygous for any of the AtPDS5 genes do not show fertility phenotypes. The functions of Arabidopsis PDS5 orthologs need to be further investigated.

1.7.1 Protection of centromeric cohesion in Arabidopsis In addition to the Sgo orthologs, AtSGO1 and AtSGO2, Arabidopsis utilizes a third factor, PANS1, for meiotic centromeric cohesion protection. AtSGO1 and AtSGO2 are utilized at anaphase I and PANS1 is required at interkinesis (Cromer et al., 2013; Zamariola et al., 2013). AtSGO1/AtSGO2 and PANS1 are all essential genes for reproduction. In Atsgo1/Atsgo2 double mutant plants, sister chromatid cohesion is lost at anaphase I and up to ten single chromatids are observed during late anaphase I, interkinesis and metaphase II. SYN1 is not detected at metaphase II in the Atsgo1/Atsgo2 double mutant. PANS1 is only found in land plants. Arabidopsis PANS1 contains a DEN box, a D box and a FLHDH motif. Both the DEN box and the D box interact with the APC/C. In Atpans1 plants, the number of chromatids is normal in late anaphase I and interkinesis, but up to ten single chromatids are observed at metaphase II. Similar

to the Atsgo1/Atsgo2 double mutant, SYN1 is not detected at metaphase II in Atpans1 mutant plants. In general, cohesion protection in Arabidopsis appears to be more complicated than in non-plant species. The mechanism of how PANS1 protects cohesin has not yet been elucidated. There are three proposed mechanisms: (1) PANS1 may protect SGO1 from Separase cleavage, (2) PANS1 may protect REC8 at centromeres, or (3) PANS1 may utilize other pathways for cohesion protection. Studies describing how cohesin is protected during mitosis in Arabidopsis have not yet been published.

1.7.2 Separase in Arabidopsis Arabidopsis contains a single copy of Separase. The predicted AtESP1 protein is much larger than ESP1 from yeast, worm or fly, but similar to mammalian ESP1. Similar to other orthologs, AtESP1 contains an EF-hand calcium-binding domain. AtESP1 also contains a C-50 peptidase domain, which is larger than those from other organisms, as it contains an extra 2Fe2S- ferredoxin domain. Studies show AtESP1 is an essential gene in Arabidopsis (Liu and Makaroff, 2006) that also functions in microtubule stability (Yang et al., 2011). Homozygous Atesp1 seeds from T-DNA heterozygous plants contain endosperm with enlarged nuclei that do not undergo cellularization and embryos that arrest at the globular stage (Liu and Makaroff, 2006). Plants containing 35S:RNAi cannot be obtained. However, plants transformed with DMC1-AtESP1- RNAi are viable and contain meiocytes that display stretched chromosomes at anaphase I and II, indicating AtESP1 is essential for the resolution of sister chromatid cohesion during meiosis I and meiosis II (Liu and Makaroff, 2006; Yang et al, 2009). Other defects including chromosome bridges and DNA fragmentation are also detected throughout meiosis (Liu and Makaroff, 2006; Yang et al, 2009). In DMC1-AtESP1-RNAi plants, SYN1 is found at the chromosome arms and centromeres during all the stages of meiosis. A temperature sensitive AtESP1 mutant has also been identified. It is named radially swollen 4 (rsw4) and contains a mis-sense mutation in AtESP1 (Wu et al, 2010). Chromosomes fail to disjoin in rsw4 roots at restrictive temperatures (Wu et al, 2010). The rsw4 roots contain higher levels of cyclin B1;1 and display disorganized cortical microtubules. Therefore, AtESP1 has functions beyond cohesin removal.

1.8 Cohesin complex: functions beyond sister chromatid cohesion Apart from its essential roles in sister chromatid cohesion, the cohesin complex plays important functions in the regulation of transcription, DNA damage repair, chromosome condensation, homolog pairing and sister kinetochore monoorientation. Some of these functions may be independent of its canonical roles in sister chromatid cohesion. In this section, the functions of cohesin in transcriptional regulation and DNA double-strand break repair will be reviewed, as well as human malignancies associated cohesin mutations.

1.8.1 Transcriptional regulation related to the cohesin complex Increasing evidence has shown that cohesin functions in transcriptional regulation. In yeast, Scc1 reduction results in gene expression changes and a reduction in DNA repair, but does not affect centromeric cohesion (Dorsett et al., 2012; Sonoda et al., 2001). The cohesin complex appears to regulate gene expression by controlling ribosomal RNA (rRNA) and protein translation in Saccharomyces cerevisiae and human (Bose et al., 2012). Cohesin also regulates the long-range interactions of enhancers and promoters as the cohesin ring entraps sister chromatin fibers and creates loops by entrapping distant chromatin segments, by interacting with CTCF or tethering the distant chromosomal loci (Wendt et al., 2008; Degner et al., 2011; Kim et al., 2011). In yeast, cohesin regulates transcriptional termination at silent mating-type loci by restricting the spread of silencing (Chang et al., 2005; Donze et al., 1999; Gullerova and Proudfoot, 2008). In mammals, cohesin colocalizes with the site-specific zinc finger DNA binding protein CTCF at some loci (Parelho et al., 2008; Rubio et al., 2008). CTCF defines the boundaries between active and inactive chromatin domains (Parelho et al., 2008; Rubio et al., 2008; Stedman et al., 2008; Wendt et al., 2008) and can recruit cohesin to specific locations. For example, both CTCF and cohesin are recruited to the imprinted H19/IGF2 locus and function in preventing the activation of the maternal IGF2 allele by an enhancer at the 3’ end of the H19 gene (Schoenherr et al., 2003). In pro B cells, cohesin is recruited to CTCF sites of V and J segments of immunoglobulin genes to facilitate gene rearrangement by contracting the locus (Degner et al., 2009). In T cells, cohesin entraps distant chromatin segments to form a loop at the IFNG locus to mediate gene expression (Mukhopadhyay et al., 2004; Hadjur et al., 2009). There are human diseases and other defects associated with cohesin mutations where sister

chromatid cohesion is unperturbed. In Cornelia de Lange Syndrome (CdLS) patients, cohesin loading is reduced and gene expression is misregulated, while sister chromatid cohesion is not severely disturbed. Cohesin also functions during nonmitotic processes as cohesin localizes within the nuclei at postmitotic stages. In Drosophila, cohesin inactivation leads to inactivation of ecdysone receptor gene expression and therefore results in pruning defects of mushroom-body γ-neurons (Lee et al., 2000).

1.8.2 Cornelia de Lange Syndrome (CdLS) Mutations in Scc2/Scc4 or components of the sister cohesin complex lead to cohesinopathy syndromes. In human, malfuntions of the cohesin complex can lead to the well-known syndromes such as Cornelia de Lange Syndrome (CdLS), Roberts Syndrome (RBS) and Warsaw Breakage Syndrome (WABS). CdLS is caused by mutations in the ATPase domains of SMC1 or SMC3 or in NIPBL, the human ortholog of SCC2 (Krantz et al., 2004; Tonkin et al., 2004). CdLS patients display various defects in growth and cognition: bone abnormalities, gastroesophageal anomalies, cardiac dysfunctions, craniofacial dysmorphia and mental retardation. Sister chromatid cohesion is not severely disturbed, but cohesin loading is defective in CdLS cells. CdLS is dominantly inherited as the disease occurs when the mutation is present in one allele of the gene. In NIPBL muations, protein levels are reduced to 70% (Krantz et al., 2004; Tonkin et al., 2004). Mutations in the ATPase domains of SMC1 or SMC3 exhibit similar phenotypes as NIPBL mutations, indicating that the association between cohesin and the chromosome is reduced (Arumugam et al., 2003; Deardorff et al., 2007; Musio et al., 2006). In CdLS cells, gene expression is also misregulated (Kawauchi et al., 2009). As demonstrated in D. melanogaster, NIPPED-B, the SCC2 ortholog, mediates long-range gene expression by interacting with regulatory sequences (Dorsett et al., 2005; Rollins et al., 1999). Mutations in Mau-2, the C. elegans Scc4 ortholog, lead to defects in axon guidance (Benard CY et al., 2004).

1.8.3 Transcriptional mis-regulation in Roberts Syndrome (RBS) The human genome contains two Eco1/Ctf7 homologs, Esco1 and Esco2 (Hou and Zou, 2005). Like the orthologs from other species, the human homologs are required for sister chromatid cohesion establishment during DNA replication. Deletion of Esco2 causes severe defects in cohesion establishment and chromosome segregation (Whenlan et al., 2012). Aneuploidy is often observed due to improper chromosome segregation. Mutations in Esco2, which is located at 8p21.1, lead to Roberts Syndrome (RBS, OMIM #268300) and SCphocomelia (OMIM #269000). Both RBS and SCphocomelia are autosomal recessive disorders. RBS was first described by Virchow in 1898 and named after John B Roberts in 1919 (Roberts, 1919). SC phocomelia Syndrome has milder phenotypes compared to RBS and was first reported by Herrmann et al. in 1969 (Herrmann et al., 1969). The symptoms of RBS patients vary significantly, indicating ESCO2 may function in a dosage-sensitive way. RBS is allelic as both severely and mildly affected individuals are observed in the same sibship. Patients with RBS show growth retardation, limb reduction/asymmetric limb growth, cleft lip/palate defects or missing fingers/toes and mild-to-severe mental deficiency (Roberts 1919; Herrmann and Opitz 1977; Van Den Berg and Francke 1993; Vega et al. 2010). RBS cells display a wide range of defects, including cohesion defects, a reduction in proliferation capacity, an increase in chromosome missegregation, cell cycle check point activation at G2/M and aneuploidy/micronuclei (Vega et al., 2005). It seems that ESCO2 is more important at the centromeres than chromosome arms, as the centromeres rather than the chromosome arms are severely affected in some cases (Williams et al., 2003). The centromeres prematurely separate. However, cohesion at the chromosome arms does display premature sister chromatid separation under DNA-damaging conditions such as mitomycin C or camptothecin treatment (Van Den Berg and Francke, 1993). The different phenotypes in RBS cells with or without DNA-damaging agents indicate ESCO2 plays distinct functions in sister chromatid cohesion establishment in mitosis and post-mitosis. In addition to the severe defects in cohesion, RBS cells also demonstrate abnormalities in gene expression, defects in nuclear morphology, and impaired ribosomal RNA (rRNA) production and protein translation (Harris et al., 2014; Lu et al., 2014). Sometimes, defects are observed in RBS cells which do not display defects in cohesion. Some RBS cells demonstrate heterochromatin repulsion (Judge 1973; Freeman et al. 1974; Tomkins et al. 1979). Cohesin can

influence the expression of genes in heterochromatin regions or alter the gene expression of adjacent euchromatin areas by controlling the spreading of heterochromatin (Ryu et al., 2014; Schamberger et al., 2014). In S. pombe, cohesin physically interacts with SWI6, the HP1 ortholog, and participates in the interaction between SWI6 and the heterochromatin (Bernard et al., 2001; Nonaka, et al., 2002). In Drosophila, cohesin binds heterochromatin regions to mediate gene expression (Vermakk and Malik, 2009). Therefore, some developmental defects in RBS/SC Syndrome may be due to altered gene expression at heterochromatin regions (Schmidt 2010). RBS cells from eco1-W216G and W539G cases exhibit defects in ribosome biogenesis and a deficit in protein translation but display no significant defects in cohesion or chromosome segregation (Harris et al., 2014; Gordillo, 2008). Meanwhile cohesion defects have been observed at the rDNA locus in an RBS patient (Bose et al., 2012). RBS cells also display apoptosis, which may be due to altered transcription of genes involved in apoptosis. Therefore, ESCO2 can regulate gene expression by affecting heterochromatin boundries or altering general translational efficiency without affecting cohesion function.

1.8.4 DNA double-strand break repair The cohesin complex also plays essential roles in DNA double-strand break repair. In response to DNA double-strand breaks in S-phase and post S-phase, the cohesin complex brings the two sister chromatids together to allow the utilization of the intact strands as a template for homologous recombination and for DNA repair. During meiosis, reciprocal recombination events create chiasmata during pachytene. Cohesin participates in the formation of the synaptonemal complex backbone (Bannister et al., 2004; Xu et al., 2005) and functions in meiotic double-strand break repair (Storlazzi et al., 2003). REC8 also mediates the repair of double-strand breaks by the use of nonsister chromatids (Bannister et al., 2004; Xu et al., 2005). ECO1/CTF7 activity is increased by DNA damage. During postreplicative DSB repair, cohesion is established by ECO1/CTF7 at the sites of damaged DNA as well as along all the chromosomes (Str¨omL et al., 2007; Unal et al., 2007; Takahashi et al., 2004). SCC1 is phosphorylated at Ser83 by checkpoint kinase1 (Chk1) and SCC1 is acetylated at K84 and K210 by ECO1/CTF7 during mitotic DNA repair. In Arabidopsis, RAD21.1/SYN2 is important for DSB repair. When Arabidopsis plants are exposed to -rays or X-rays, SYN2 expression level is increased (da Costa-Nunes et al., 2006;

Kozak et al., 2009). However, no vegetative phenotype is observed in syn2 plants when treated with X-rays or bleomycin. Recent studies demonstrate that Arabidopsis WAPL and CTF7 also play essential roles in DNA double strand break repair during mitosis (De, Bolaños-Villegas and Makaroff, unpublished).

1.8.5 Human cancers related to mis-function of cohesin Cohesin and cohesion-associated proteins are associated with several human cancers. In colorectal cancers, NIPBL, SMC1A, SMC3, RAD21, SA2 and STAGE3 are frequently mutated (Barber and McManus, 2008). SMC1A and SMC3 are important for the supression of chromosome instabilities (CIN) and mutations in the ATPase domains of SMC1 or SMC3 lead to various defects in cohesin’s association with chromosomes (Arumugam et al., 2003; Arumugam et al., 2006). In acute myeloid leukemia, SMC1A, SMC3, RAD21 and SA2 are often mutated, even though aneuploidy and CIN do not occur (Welch et al. 2012). In human melanoma cells, ESCO2 is overexpressed (Ono et al., 2003). In cervical cancers, WAPL expression is increased and reducing expression of WAPL lessens the severity of cervical cancers (Oikawa et al., 2004). Separase functions as an oncogene in tumorigenesis (Zhang et al., 2008b). In epithelial tumors, heterozygous separase mutation leads to genome instability and initiates cancer (Zhang et al., 2008b). In addition, separase overexpression induces aneuploidy and tumorigenesis in mammary epithelial cells of mouse if p53 is mutated (Pati et al., 2004; Zhang et al., 2008b). In human cells, mutations in BRCA1 cause similar defects as mutations in cohesin complex subunits (Kobayashi et al., 2004; Mayer et al., 2004; Petronczki et al., 2004). BRCA1 interacts with cohesin complex subunits and the functions of cohesin and BRCA1 overlap in tumorigenesis. In zebrafish, RAD21 regulates the transcription of MYCA, p53 and Mdm2, and their mis-regulation is related to cancers (Mönnich et al., 2011). Heterozygous SA1 mice display increased aneuploidy and tumorgeneisis (Remeseiro, Cuadrado et al. 2012). Even though the association of cohesin dysfunctions and cancer development are clear, the relationship between them needs to be further elucidated. In different cohesinopathy diseases, similarities in gene expression changes and disease phenotypes are observed. The similarities are caused by a deficiency in the cohesin subunits which are essential for cohesion. Meanwhile, differences are also observed in different cohesionpathies as different components play different roles. CdLS and RBS exhibit distinct

phenotypes as they are caused by mutations in different subunits of the cohesin complex (Liu and Krantz, 2009; Nasmyth and Haering, 2009).

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Chapter 2: 35S:NTAP:AtCTF7∆B leads to various defects in reproduction

2.0 Abstract

Eco1/Ctf7 establishes sister chromatid cohesion during meiosis and mitosis. Eco1/Ctf7 is an essential gene in many species. In human, mutations in Esco2, the Eco1/Ctf7 ortholog, lead to Roberts Syndrome (RBS), which results in various defects in cohesion and other processes. Inactivation of Arabidopsis CTF7 (AtCTF7) by T-DNA mutation or reducing AtCTF7 expression levels by RNAi leads to severe defects in reproduction and vegetative growth. To further investigate the potential roles of AtCTF7 and to identify AtCTF7 interacting proteins, several AtCTF7 constructs were generated and expressed in Arabidopsis plants. Among them,

35S:NTAP:AtCTF7∆B (AtCTF7∆299-345) transgenic plants displayed a wide range of phenotypic alterations in reproduction. Male meiocytes from 35S:NTAP:AtCTF7∆B plants exhibited chromosome fragmentation and defective chromosome segregation. Mutant ovules developed asynchronously and experienced elongated meiosis and mitosis. The mutant ovules also produced degenerated megaspores/embryo sacs at various stages as well as megaspores with altered identities. Previous studies showed that mutations in Eco1/Ctf7 lead to defects in rRNA production and protein biosynthesis (Guacci et al., 1997; Gard et al., 2009; Heidinger-Pauli et al., 2010). Similar defects between 35S:NTAP:AtCTF7∆B ovules and cells with mutations in RNA processing genes suggest that RNA processing may be disrupted in 35S:NTAP:AtCTF7∆B plants.

This chapter along with chapter 3 has been submitted to Plant Physiol.

2.1 Introduction

2.1.1 Background for ECO1/CTF7 Sister chromatid cohesion is established during S phase by ECO1/CTF7. ECO1/CTF7 exhibits acetyltransferase activity (Ivanov et al., 2002; Bellow et al., 2003). During cohesion establishment, ECO1/CTF7 acetylates conserved lysine residues of SMC3, which are close to the ATP binding domain (Rowland et al., 2009; Zhang et al., 2008a). ECO1/CTF7 also acetylates lysine residues of SCC1 and other proteins (Ivanov et al., 2002; Ghosha et al., 2012). Acetylated SMC3 stabilizes the interaction between SMC3 and SCC1, and counteracts WAPL removal of cohesin from the chromosomes, allowing sister chromatid cohesion to be established (Ben- Shahar et al., 2008; Unal et al., 2008; Zhang et al., 2008a; Rowland et al., 2009; Sutani et al., 2009). In budding yeast, ECO1/CTF7 acetylates conserved lysine residues, K112 and K113, in SMC3 to establish sister chromosome cohesion (Rowland et al., 2009). In the eukaryotes, the corresponding residues (K112 and K113) of SMC3 are conserved and acetylated (Rowland et al., 2009). In many species, ECO1/CTF7 is an essential gene and inactivation of Eco1/Ctf7 leads to various defects such as chromosome mis-organization, cohesin protein mis-distribution, cell cycle check point activation and growth retardation (Milutinovich et al., 2007, Skibbens et al., 1999, Toth et al. 1999). In addition to the essential roles in cohesion establishment, Eco1/CTF7 is also important for nucleolar integrity, rRNA production, ribosome biogenesis and protein biosynthesis (Guacci et al., 1997; Gard et al., 2009; Heidinger-Pauli et al., 2010). Cohesion establishment occurs in S phase and is coupled to DNA replication. ECO1/CTF7 interacts with various factors: DNA replication factors (Mayer et al., 2001), CTF4 and CTF18 (Lengronne et al., 2004), PCNA (DNA polymerase processivity factor) (Moldovan et al, 2006) and the DNA helix (Mayer et al., 2004). Interestingly, deletion of Rad61/Wapl or mutations in specific domains of Pds5, Smc3 and Scc3 rescue the Eco1/Ctf7 deletion, suggesting that Eco1/Ctf7 cohesion establishment activity is dispensible in the absence of the anti-establishment genes (Ben-Shahar et al., 2008; Rowland et al., 2009; Sutani et al., 2009), and that is acetylation of Smc3 is not required without the

“antiestablishment” activity. Alternatively, the modified cohesin may counteract the Eco1/Ctf7 cohesion anti-establishment activity (Sutani et al., 2009; Ben-Shahar et al., 2008; Heidinger- Pauli et al., 2008; Rowland et al., 2009; Unal et al., 2008; Zhang et al., 2008a; Shintomi and Hirano, 2009).

2.1.2 Esco2 mutations in Roberts Syndrome (RBS) There are two Eco1/Ctf7 homologs, Esco1 and Esco2, in the human genome (Hou and Zou, 2005). Mutations in Esco2 lead to Roberts Syndrome (RBS). Cells from RBS patients typically contain Esco2 deletions or insertions, which result in frameshifts (61.5%), splice-site mutation (23.1%), nonsense (11.6%) or missense (3.8%) mutations (Vega H et al., 2010; Gordillo et al., 2008). Most Esco2 mutations lead to premature stop codons prior to or within the acetyltransferase domain, which lead to complete or partial loss of the acetyltransferase activity. Therefore, acetyltransferase activity is essential for ESCO2 function (Gordillo et al., 2008). RBS cells exhibit a wide range of defects, including cohesion defects, a reduction in proliferation capacity, defects in nuclear morphology, abnormalities in gene expression, and impaired ribosomal RNA (rRNA) production and protein translation. In some RBS cells, cohesion is unperturbed but gene expression patterns are altered (Harris et al., 2014; Lu et al., 2014)

2.1.3 The AtCTF7 protein The Arabidopsis genome contains one Eco1/Ctf7 ortholog. AtCTF7 plays essential roles in Arabidopsis and AtCTF7 is able to rescue the yeast temperature-sensitive mutant ctf7-203 (Jiang et al., 2010). AtCTF7 is a 345 amino acid protein that contains a PIP box (QxxL/I, QFHL) at residues 82 to 86, a C2H2 zinc finger motif at residues 92 to 130 and an acetyltransferase domain at residues 184 and 335. The acetyltransferase domain can be further separated into three motifs: D (184-204aa), A (266-301aa) and B (311-335aa) (Figure 2.2A). Motif D provides the framework of the domain. The A motif participates in Acetyl CoA binding and is critical for catalytic activity. The B motif is the C-terminal most region; it participates in substrate recognition and the regulation of catalytic activity (Jiang et al., 2010; Ivanov et al., 2002). Similar to S. cerevisiae ECO1, AtCTF7 does not contain a large extension at N terminus. ECO1/CTF7 sequences are conserved among the eukaryotes with the proteins sharing significant similarity (Table 2.1, Figure 2.1). AtCTF7 exhibits 24% to 58% similarity with

ECO1/CTF7 from yeast, fruit fly and vertebrates. AtCTF7 shows the highest sequence similarity to human ESCO1 (22% identity, 58% similarity).

Table 2.1 ECO1/CTF7 orthologs (Yuan et al., 2011).

A. thaliana S.cerevisiae S.pombe D.melan Vertebrates Rice Maize

ogaster CTF7 ECO1/ ESO1 DECO ESCO1 OSI_19739 ESCO1 CTF7 ESCO2

Figure 2.1 ECO1/CTF7 sequences are conserved among eukaryotes.

The PIP box, C2H2 zinc finger motif are marked as well as the three motifs (D, A and B) of the acetyltransferase domain. Identical residues are marked as * and similar residues are marked as : .

ECO1/CTF7 acetylates the cohesin subunits, SMC3, SCC1, SCC3 and PDS5, but not histones in vitro (Ivanov et al., 2002; Ghosha et al., 2012). Its zinc finger motif regulates its functions and increases the acetyltransferase activity (Zou et al., 1999; Ivanov et al., 2002). Just like orthologs from other species, AtCTF7 displays acetyltransferase activity in vitro (Jiang et al., 2010). When AtCTF7 was fused with the maltose-binding protein (MBP), overexpressed in Escherichia coli and purified, the protein demonstrated autoacetylation activity (Jiang et al., 2010). Much progress has made on understanding ECO1/CTF7 functions; however no ECO1/CTF7 structure has been solved. By bioinformatic methods, the AtCTF7 structure has been proposed based on structures of N-acetyltransferase family proteins (Figure 2.2B) (Zhang, 2008b; Roy et al., 2010; Roy et al, 2012). From the proposed structure, the N terminus of AtCTF7 is random while the C terminal acetyltransferase domain is well folded. The random N terminus shows its flexibility in factor binding and regulatory functions for acetylation activity. The C terminal acetyltransferase domain contains an acetyl-coenzyme A (Acetyl CoA) binding site that belongs to the GNAT (Gcn5-related N-acetyltransferase) family (Ivanov et al., 2002). The three motifs of the acetyltransferase domain play different roles: motif D provides the framework for the protein, while motif A helps AcCoA binding and is critical for the catalytic activity. Motif B is the most C-terminal region, participating in substrate recognition and activity regulation.

(A)

MQAKINSFFKPSSSSSIAASVTTDTDDGLAVWENNRNAIVNTYQRRSAITERSEVLKGCIE KTLKKGSSSVPKNHKKKRNYTQFHLELGQSDFLLRHCAECGAKYAPGDELDEKNHQSF HKDYMYGLPFKGWQNEKAFTSPLFIKNRIVMVSENDSPAHRNKVQEVVKMMEVELGE DWILHQHCKVYLFISSQRISGCLVAEPIKEAFKLIASPDDERQLQKESSSSPSTSIQFGNIVL QREVSKRCRTSDDRLDNGVIVCEEEAKPAVCGIRAIWVSPSNRRKGIATWLLDTTRESFC NNGCMLEKSQLAFSQPSSIGRSFGSKYFGTCSFLLYKAQLIDTHFS (B)

Figure 2.2 The amino acid sequence of AtCTF7 and the proposed structure of AtCTF7. (A) The amino acid sequence of AtCTF7.

AtCTF7 is 345 amino acids long and contains a PIP box (blue) at residues 82 to 86, a C2H2 zinc finger motif (green) at residues 92 to 130 and an acetyltransferase domain from residues 184 to 335 (red). (B) The proposed structure for AtCTF7. AtCTF7 belongs to the GCN5-related N-acetyltransferase superfamily. The N terminus is predicted to have a random structure while the C terminal acetyltransferase is well folded. The model was generated using I-TASSER server (Zhang, 2008b; Roy et al., 2010; Roy et al, 2012).

Functions of CTF7 in Arabidopsis Two T-DNA insertion lines, SALK_059500 (ctf7-1) and SAIL_1214G06 (ctf7-2), have been characterized (Jiang et al., 2010; Bolaños-Villegas et al., 2013). The T-DNA insertions in ctf7-1 and ctf7-2 are in intron 4 and exon 3, respectively. Just like RBS cases, the T-DNA insertions disrupt the protein upstream of the acetyltransferase domain. While heterozygous Atctf7+/- plants are normal in vegetative growth and pollen development, homozygous Atctf7-/- plants are severely dwarf and sterile (Bolaños-Villegas et al., 2013; Jiang et al., 2010). Quantitative real- time PCR (qPCR) revealed that homozygous Atctf7 plants produce about 20% of normal AtCTF7 mRNA levels after the T-DNA insertion sites and produce normal levels of AtCTF7 mRNA before the T-DNA insertions. Therefore, in both Atctf7-1 and Atctf7-2 plants, truncated versions of the transcripts and possibly truncated proteins are produced, resembling the RBS mutations (Vega et al., 2005). In addition to the vegetative growth defects, fertility defects are observed (Bolaños-Villegas et al., 2013; Jiang et al., 2010). Even though heterozygous Atctf7+/- plants show normal vegetative growth, their siliques produce fewer seeds (46 seeds/silique) and approximately 25% of the seeds are aborted after fertilization (Bolaños-Villegas et al., 2013; Jiang et al., 2010). Unexpectedly, homozygous Atctf7 mutant plants can be detected at a very low frequency, about 4% of the seeds from heterozygous Atctf+/- plants. The siliques from Atctf7-/- plants also produce fewer ovules (22 ovules/silique) and the ovules that are produced are unfertilized and abort early. Knockdown of AtCTF7 mRNA levels using RNAi also leads to growth retardation (Singh et al., 2013). Finally, overexpression of full length AtCTF7 using the 35S promoter (35S:AtCTF7) leads to reduced fertility of varying degrees, but does not afftect pollen development or vegetative growth (Jiang et al., 2010). In general, AtCTF7 is required for ovule/seed development from ovule germination until seed maturation. In heterozygous Atctf+/- plants and in Atctf7-/- plants, the Atctf7- ovules/seeds are dramatically different, which might be due to different amounts of residual AtCTF7 mRNA (Jiang et al., 2010; Bolaños-Villegas et al., 2013). Or the surrounding somatic cells may help mediate Atctf7 embryo sac development by providing AtCTF7 mRNA/protein, similar to the situation for AGO9 (Olmedo-Monfil et al., 2010)

2.2 Results

2.2.1 Experimental design and dominant negative defects of 35S:NTAP:AtCTF7∆B plants To further investigate potential roles of AtCTF7 and to identify AtCTF7 interacting proteins, a 35S:NTAP:AtCTF7∆B construct was generated and transformed into wild type plants. The 35S:NTAP:AtCTF7∆B construct is missing the C-terminal 46 amino acids (∆299-345), which corresponds to the acetyltransferase B motif, and is expressed from the CaMV 35S promoter (Figure 2.3). Columbia plants transformed with 35S:NTAP:AtCTF7∆B displayed various alterations in vegetative growth and reduced fertility.

Figure 2.3 The 35S:NTAP:AtCTF7∆B construct. The 35S:NTAP:AtCTF7∆B construct is missing the C-terminal 46 amino acids of the acetyltransferase B motif (∆299-345). It is expressed from the CaMV 35S promoter. The TAP tag contains calmodulin binding peptide (CBP) at the N-terminal, followed by tobacco etch virus protease cleavage site cleavage site (TEV protease site) and Protein A site.

Twenty out of the 36 independent 35S:NTAP:AtCTF7∆B lines examined exhibited reduced fertility, with the fertility levels varying significantly between the lines. Lines exhibiting a weak phenotype, which accounted for two of the 20 reduced fertility lines, produced shorter siliques with fewer seeds, but the seeds looked normal (Figure 2.4Aii). For example Line 19 produced 39.2±3.9 seeds per silique (n =35) compared to wild type siliques that produce 54.2±4.1 seeds per silique (n=35). Anthers from Line 19 plants were smaller but contained fully viable pollen (Figure 2.4Bii). The number of pollen in line 19 was approximately half compared to wild type (574 verses 1175). The other 18 lines exhibited more severe defects. The plants produced siliques containing large numbers of unfertilized ovules and aborted seeds (Figure 2.4Aiii). Seed set

varied considerably between the lines, ranging from 13.3±5.6 seeds per silique to 34.7±8.4 seeds per silique. Unfertilized ovules appeared as white dots, resembling the situation in Atctf7-1 plants (Bolaños-Villegas et al., 2013). The aborted seeds appeared white and plump, similar to seeds containing arrested embryos in Atctf7-1+/- plants (Jiang et al., 2010). The different numbers of defective ovules/seeds in the same silique and large range of seed set in different lines suggested that the expression of the 35S promoter was variable, that different pathways may be disrupted or possibly that the defects were cumulative. Anthers from the severely reduced fertility plants typically contained few-viable pollen (210 in line 11, Figure 2.4Biii). Given that most lines exhibited severe fertility defects, one representative line (#11) was chosen and characterized in detail, unless specified otherwise.

Figure 2.4 35S:NTAP:AtCTF7∆B plants exhibit reduced fertility. (A) Seed set in wild type Columbia plants (i) and in 35S:NTAP:AtCTF7∆B plants exhibiting mild (ii) and severe (iii) fertility defects. (i) Wild type silique with full seed set. (ii) Silique from a 35S:NTAP:AtCTF7∆B plant exhibiting a weak phenotype (Line 19). All seeds look normal. (iii) Silique from a 35S:NTAP:AtCTF7∆B plant exhibiting a strong phenotype (Line 11). Arrows indicate shriveled, unfertilized ovules. White stars show white, plump seeds, which are aborted after fertilization. Scale bar = 0.5 cm. (B) Alexander staining of mature anthers from wild type (i) and 35S:NTAP:AtCTF7∆B plants (ii, iii). (i) Mature wild type anther full of viable pollen. (ii) Mature anther from Line 19. The anther is smaller and contains less pollen; all the pollen is viable. (iii) An anther from Line 11. The anther is smaller and contains low numbers of viable pollen (42 viable pollen vs 210 total pollen). Scale bar =50 µm.

Preliminary analysis of 35S:NTAP:AtCTF7∆B plants indicated that both male and female fertility are affected in the lines. Reciprocal crossing experiments showed that the 35S:NTAP:AtCTF7∆B construct was transmitted at reduced levels through both the male and female gametes (Table 2.2). Nonviable seeds were obtained when 35S:NTAP:AtCTF7∆B plants were used as either the male or female parent; however the relative proportion of nonviable seeds was three times greater (9.3% verses 31.3%) when 35S:NTAP:AtCTF7∆B plants served as the female. The number of nonviable seeds was greatest (65%) in self-pollenated plants, indicating there is an additive effect.

Table 2.2 35S:NTAP:AtCTF7∆B transfer efficiency Female (♀) X Seeds that did not Basta Resistant Basta Sensitive male(♂) germinate

WT X Line 11 137 (38.3%) 188 (52.5%) 33 (9.3%)

Line 11 XWT 71 (25%) 124 (43.7%) 89 (31.3%)

Line 11 self-pollinated 354 (30.4%) 49 (4.2%) 763 (65.4%)

4th generation Line 11 plants were used in the cross. If a plant is Basta resistant, it contains the transgene 35S:NTAP:AtCTF7∆B construct. If a plant is Basta sensitive, it does not contain the transgene.

The reduced fertility phenotype of 35S:NTAP:AtCTF7∆B plants suggested that co-suppression could be occurring to reduce AtCTF7 transcript levels in the lines. Therefore, quantitative reverse transcription polymerase chain reaction (qRT-PCR) was conducted to examine transcript levels of native AtCTF7 as well as the AtCTF7 transgene (Figure 2.5). In severely reduced fertile

transgenic plants (Line 11), total AtCTF7 (35S:NTAP:AtCTF7∆B and native AtCTF7) transcript levels were about 7 fold higher than AtCTF7 transcript levels in wild type plants, while native AtCTF7 transcript levels were almost 2.8 fold higher in transgenic plants. In mildly reduced fertility plants (Line 19), total AtCTF7 transcript levels were about 5.4 fold higher than wild type and native AtCTF7 transcript levels were 1.4 fold higher. Therefore, the transgenic lines contained high levels of AtCTF7∆B transcripts and elevated levels of native AtCTF7 mRNA, indicating that the observed phenotypes were not the result of reduced AtCTF7 expression. Instead it suggests that the 35S:NTAP:AtCTF7∆B construct is exerting a dominant negative effect. The varying severity in different 35S:NTAP:AtCTF7∆B lines might be due to different levels of AtCTF7.

Figure 2.5 Expression analysis of AtCTF7 in wild-type and 35S:NTAP:AtCTF7∆B plants. Transcript levels of total AtCTF7 (from 35S:NTAP:AtCTF7∆B and from native AtCTF7) were about 7-fold higher and transcript levels of native AtCTF7 were about 2.8-fold higher in Line 11. Transcript levels of total AtCTF7 were about 5.4-fold higher and transcript levels of native AtCTF7 were about 1.4-fold higher in Line 19. Buds of WT, non-dwarf 35S:NTAP:AtCTF7∆B plants (4th generation of Line 11) and 4th generation plants from Line 19 are used. Tublin was used as the internal control.

2.2.2 35S:NTAP:AtCTF7∆B plants demonstrate defective chromosome segregation during male meiosis Previous results showed that inactivation of AtCTF7 via T-DNA insertion results in uneven chromosome segregation during meiosis and ultimately reduced pollen viability (Bolaños- Villegas et al., 2013). The cohesin protein SYN1 distribution is also altered in homozygous ctf7- 1 male meiocytes. The reduced fertility observed in 35S:NTAP:AtCTF7∆B plants suggested that meiosis might be affected. Therefore, chromosome spreading experiments were carried out to test the effect of 35S:NTAP:AtCTF7∆B expression on male meiosis. Male meiocytes in 35S:NTAP:AtCTF7∆B plants resembled wild-type during early stages of meiosis, with normal chromosome morphology during pachytene (Figure 2.6A,E), diakinesis (Figure 2.6B,F) and metaphase I (Figure 2.6C,G). The first noticeable defect was fragmented chromosomes, which were observed at anaphase I ((Figure 2.6H). Mis-segregated chromosomes were observed at prophase II (Figure 2.6M). More severe segregation defects were detected during meiosis II. At metaphase II, the meiocytes contained more than 10 chromosomes (Figure 2.6N), indicating that cohesion was lost. The chromosomes did not segregate evenly at anaphase II (Figure 2.6O) with the production of polyads with varying DNA content (Figure 2.6P). Similar to Atctf7-/- (Bolaños-Villegas et al., 2013), some relatively normal meiocytes were also observed throughout the pollen development.

Figure 2.6 35S:NTAP:AtCTF7∆B meiocytes demonstrate defective meiotic chromosome organization. DAPI-stained male meiotic chromosomes from the WT (A–D, I-L) and 35S:NTAP:AtCTF7∆B meiocytes (E-H, M-P). A and E, pachytene. B and F, diakinesis. C and G, metaphase I. D and H, anaphase I. I and M, prophase II. J and N, metaphase II. K and O, anaphase II. L and P, telophase II. Lagging chromosomes or chromosome fragments are denoted with arrows. Scale bar =10 μm.

Immunolocalization experiments were then carried out to examine the distribution of the meiotic cohesion protein SYN1 on 35S:NTAP:AtCTF7∆B male meiocytes (Figure 2.7). The normal distribution of SYN1 protein implied that SYN1 protein levels and their distribution were not affected or the change was not dramatic enough to be revealed by immunolocalization experiments.

Figure 2.7 Cohesion protein SYN1 distribution pattern is not affected in 35S:NTAP:AtCTF7∆B male meiocytes. Male meiocytes of WT (A-C) and 35S:NTAP:AtCTF7∆B plants (D-F). Interphase/early leptotene (A,D); zygotene (B,E), pachytene (C,F). Merged images of 4′,6-diamidino-2-phenylindole (DAPI) stained chromosomes (red) and SYN1 (green) are shown. Scale bar =10 μm.

2.2.3 35S:NTAP:AtCTF7∆B leads to various defects in ovule development

2.2.3.1 Ovule development in wild type Arabidopsis Ovule development has been well defeine in Arabidopsis (Schneitz et al. 1995; Christensen et al. 1997, 1998). In WT siliques, archesporial cells are specified from the subepidermal cell layer and differentiate into the MMCs (megaspore mother cells) (Figure 2.8A). The initial MMC is un- polarized and becomes polarized prior to the start of meiosis. Megaspore development consists of two phases: megasporogenesis (meiosis, phase I) and megagametogenesis (mitosis, phase II). During megasporogenesis, the MMC undergoes two rounds of meiosis to produce a tetrad of four haploid megaspores (Figure 2.8B-E). When the ovule reaches FG1 stage, the megaspore at the chalazal-end differentiates into the functional megaspore (FM) while the other three megaspores undergo programmed cell death (Figure 2.8F,G; Figure 2.9B,C). At FG1, the outer integument, but not the inner integument encloses the nucellus. During megagametogenesis, the FM undergoes three rounds of mitosis accompanied by nuclear migration, nuclear fusion, nuclear degeneration and cellularization to form the final embryo sac (Figure 2.8H-L; Figure 2.9D-J). The FM at FG1 undergoes the 1st mitosis to produce a two-nucleate embryo sac (FG2, Figure 2.8H), followed by the formation of a vacuole between the two nuclei (FG3, Figure 2.8I, Figure 2.9D). At FG3, the ovule becomes curved and the inner integument embraces the nucellus. The gametophyte then undergoes a second mitosis to produce a four-nucleate embryo sac (FG4, Figure 2.8J, Figure 2.9E), followed by the migration of the two chalazal nuclei from the orthogonal direction to the chalazal-micropylar axis (Figure 2.9F). After nuclear migration, the third mitosis takes place and gives rise to eight nuclei in a 4n+4n configuration (FG5, Figure 2.9G). The two polar nuclei, one from each side, meet at the embryo sac’s micropylar half and fuse to become the central nucleus (CN), and the antipodal nuclei (ANs) start to degenerate (FG6, Figure 2.9H). At FG7, the two polar nuclei fuse and the ANs degenerate completely (Figure 2.9I). Prior to fertilization, one synergid nucleus (SN) degenerates and the embryo sac consists of one egg nucleus, one central nucleus and one synergid nucleus (FG8, Figure 2.8L, Figure 2.9J). Double fertilization then occurs as the egg nucleus fertilizes with one sperm to form the embryo and the central nucleus fertilizes with a second sperm to form the endosperm.

Figure 2.8 Female gametophyte development in WT Arabidopsis plants revealed by differential interference contrast (DIC) microscopy. (A) The pre-meiotic ovule at stage 1-II with elongated protrusions. The megaspore mother cell (MMC) is denoted by an arrow. (B) The meiotic ovule at stage 2-IV contains two longitudinal, divided megaspore cells after meiosis I. (C) The meiotic ovule at stage 2-IV contains two megaspore cells after meiosis I, and the bottom one contains two nuclei. (D) The meiotic ovule at stage 2-V contains a tetrad on the same plane, after meiosis II. (E) The meiotic ovule at stage 2-V contains a linear tetrad. (F) The ovule of stage 3-I at the early FG1 stage. (G) The ovule of stage 3-I at the FG1 stage. The FM (arrow) is uni-nucleate and there is still space at the distal end of the ovule. (H) The ovule of stage 3-II at the FG2 stage. The embryo sac contains two divided nuclei

(arrows). (I) The ovule of stage 3-III at the FG3 stage. A vacuole germinates between the two nuclei. (J) The ovule of stage 3-IV at the FG4 stage. (K) The ovule of stage 3-V at the FG5-6 stage. (L) The ovule of stage 3-VI at the FG7-8 stage. The embryo sac contains the egg nucleus (EN) and the central nucleus (CN). CN, central nucleus; EN, egg nucleus; MMC, megaspore mother cell; SN, synergid nucleus; V, vacuole. Scale bar=10 μm. Ovule stages are determined according to Schneitz et al. 1995.

Figure 2.9 Female gametophyte development in WT Arabidopsis plants, revealed by confocal laser scanning microscopy (CLSM). (A) FG0-stage ovule. The MMC is surrounded by the nucellar epidermis. The outer and inner integuments just start to initiate. (B) Early FG1-stage ovule, showing the FM and DM. The nucellus is not surrounded by the integuments. (C) FG1-stage ovule. The nucellus is surrounded by the outer integuments but not the inner integuments (marked as the hole). (D) FG3-stage ovule. The embryo sac consists of two nuclei separated by a central vacuole. The nucellus is enclosed by the inner integument. (E) Early FG4-stage ovule. The embryo sac contains four nuclei. The division planes of the two nuclei are orthogonal to each other. (F) Late FG4-stage ovule. A vacuole appears at the chalazal end of the embryo sac. (G) FG5-stage ovule. The embryo sac contains eight nuclei in a 4n+4n configuration. Not all the nuclei are visible on this plane. (H) FG6-stage ovule after the cellularization. The three ANs are degenerating (arrow) and the CN does not form yet. (I) FG7-stage ovule with a four-nuclei embryo sac. The CN forms and the ANs have degenerated. (J) FG8-stage ovule, just prior to fertilization. The embryo sac consists of the EN and the CN. AN, antipodal nucleus; CN, central nucleus; DM, degenerated megaspore; EN, egg nucleus; FM, functional megaspore; MMC, megaspore mother cell; SN, synergid nucleus; V, vacuole.

Bar scale=10 μm. The developmental stages are defined according to Christensen et al. 1997.

2.2.3.2 Ovules display various defects due to variations in AtCTF7 expression Previous studies showed that AtCTF7 is essential for ovule development as various defects are observed when AtCTF7 levels are disturbed (Jiang et al., 2010, Bolaños-Villegas et al., 2013). Ovules from Atctf7-1+/- heterozygous plants contain non-degenerating antipodal nuclei (Jiang et al., 2010) and the ovules from the Atctf7-/- plants degenerate early (Bolaños-Villegas et al., 2013). Decreasing AtCTF7 transcript levels by RNAi or increasing AtCTF7 transcripts by 35S:AtCTF7 arrests ovules at FG1 stage (Singh et al., 2013; Jiang et al., 2010). In Atctf+/- plants and in Atctf7-/- plants, the Atctf7 ovules have completely different fates. Atctf7- ovules from Atctf7+/- heterozygous plants contain minor defects in antipodal nuclei degeneration and the ovules can be fertilized by Atctf7- pollen. Atctf7- ovules from Atctf7-/- plants are aborted before the fertilization. The different fates of Atctf7- ovules from these plants indicate the Atctf7- ovule development is not solely controlled by the genetic background of the gametophyte/embyo sac. The different Atctf7- ovules fates could be due to carryover of AtCTF7 mRNA and/or protein. Alternatively, the surrounding somatic cells may help mediate the Atctf7- ovule development.

2.2.3.3 35S:NTAP:AtCTF7∆B leads to various defects in ovule development The effects in 35S:NTAP:AtCTF7∆B on ovule development were analyzed by differential interference contrast (DIC) microscopy and confocal laser scanning microscopy (CLSM).

35S:NTAP:AtCTF7∆B leads to asynchronous ovules development in the same silique In WT siliques, ovules grow in opposite directions at similar rates (Figure 2.10A,C). Conversely, some 35S:NTAP:AtCTF7∆B ovules developed unevenly. In Figure 2.10B, the neighboring ovules are different in size and grow in opposite directions. In Figure 2.10D, the neighboring ovules are different in size and pointing in the same direction, which was never observed in WT ovules. Ovules of different sizes might have differientated at different times or may have grown at different speeds. The ovules facing the same direction likely differientated at different times, suggesting that 35S:NTAP:AtCTF7∆B disrupted ovule development mechanisms. Beside the relatively normal MMC, there were abnormal megaspores in the ovules. Even though the mutant ovules were different in size and stage, primordial initiation and the inner and outer

integuments development were normal.

Figure 2.10 Asynchronous development of early 35S:NTAP:AtCTF7∆B ovules. (A and C) Ovules from wild type siliques. The ovules are similar in size and stage, and point in opposite directions. (B and D) Ovules from the 35S:NTAP:AtCTF7∆B siliques. (B) The ovules are different in size and grow in the opposite directions. (D) The ovules are different in size and stages (stage 1-II and stage 2-III), and point in the same direction. Abnormal megaspores (arrows) are also observed on the ovules. The morphology of ovules in B and D are normal compared to the corresponding wild type ovule. Scale bar is 10 μm.

Asynchronous and elongated ovule development in 35S:NTAP:AtCTF7∆B plants In CLSM experiments, pistils from the same inflorescence were dissected and the ovule stages were recorded accordingly to Christensen et al., 1997. In WT plants, the ovules developed synchronously with one predominate stage in each pistil (Table 2.3) (Christensen et al., 1997, 2002). Meanwhile, 35S:NTAP:AtCTF7∆B ovules display slowed and asynchronous development (Table 2.3). Female gametophytes from the same pistil were spread out in several stages with many embryo sacs arrested at FG2/FG3 stages, suggesting that the synchrony of gametophyte development was disturbed and embryo sac maturation was delayed.

Table 2.3 Female gametophyte development in WT (Col) plants Pistil # FG1 FG2 FG3 FG4 FG5 FG6 FG7 FG8 1 30 2 1 2 26 3 4 23 4 18 13 5 8 26 1 6 1 7 38 7 1 7 16 15 8 1 42

Table 2.4 Female gametophytes in 35S:NTAP:AtCTF7∆B plants

Pistil# FG0- FG0- FG1 FG1 FG2 FG- FG3 FG4 FG5 FG6 FG7 FG- FG- A B (early) (late) C D E 1 17 5 2 13 12 1 3 7 15 5 1 1 4 1 14 9 1 2 5 2 6 14 4 3 6 1 13 12 1 7 4 16 2 6 8 4 15 2 6 9 2 4 20 10 1 16 1 2 3 11 1 17 2 8 12 3 10 3 13 13 12 3 1 18 14 8 1 2 20 15 4 12 15

A fourth generation Line 11 plant was used. Female gametophyte stages were defined according to Christensenet al., 1998. FG0-A: ovules with no megaspore (Figure 2.11A); FG0-B: ovules with megaspore(s) but at the pre-meiotic stage (Figure 2.11B); FG-C: small ovules with no observable nuclei (Figure 2.13D); FG-D: final ovules containing embryo sacs (Figure 2.13N,O,P). FG-E: Final ovules without an embyso sac (Figure 2.13M).

Because gametophyte development was slow, the megaspore commitment process was easy to identify in the majority of ovules (Figure 2.11). During this process, a separated megaspore migrated to the chalazal end and became functional megaspore-like (FML) while the remaining megaspore(s) degraded (Figure 2.11B-D). Then, the FML underwent nuclear division to produce

two nuclei and a central vacuole formed between them (Figure 2.11E).

Figure 2.11 Female gametophyte development is slowed in 35S:NTAP:AtCTF7∆B plants, revealed by confocal laser scanning microscopy (CLSM). (A) No megaspore(s) is identified. (B) The ovule contains only megaspore(s). (C) A functional megaspore-like (FML) separates from the megaspores and the remaining megaspores become DM(s). (D) The FML migrates to the chalazal end. (E) The FML completes one round of mitosis to produce two nuclei and a vacuole forms between the two nuclei. Note: FM(L)s are identified as having distinctly bright autofluorescence in the nuclei and the DMs contain a diffuse signal throughout the cell but no clearly defined nucleus (Barrell and Grossniklaus, 2005).

The defects we observed in 35S:NTAP:AtCTF7∆B ovules were further examined by DIC. Consistent with CLSM results, the slowed and asynchronous gametophyte development was revealed as the ovules elongated during meiosis (Figure 2.12 A-D) and mitosis (Figure 2.12 E-G). During meiosis, various numbers of megaspores were observed in the middle of the ovules (Figure 2.12B,C). The megaspores contained apparent nuclei with variable morphologies. Based on the number of megaspores, the ovule in Figure 2.12B appears to contain an undivided MMC and the ovule in Figure 2.12C contains a dyad after meiosis I. Most gametophytes progressed beyond meiosis and arrested at FG2 (Figure 2.12F) or FG3 (Figure 2.12G), even though the overall ovule size and shape reached a later stage.

Figure 2.12 Meiosis and mitosis during female gametophyte development is slowed in 35S:NTAP:AtCTF7∆B ovules revealed by DIC. (A) The ovule contains one megaspore with an apparent nucleus (arrow); the integuments do not enclose the nucellus. (B) The ovule contains one megaspore with an apparent nucleus (arrow); the integuments completely enclose the nucellus. The megaspore is an undifferentiated MMC. (B’) Enlarged view of the MMC from E. (C) Ovule contains two megaspores with prominent nuclei (arrows). They are at the dyad stage after meiosis I. The two megaspores are clearly separated. (C’) Enlarged view of the dyad from C. (D) Ovule contains two megaspores, which are mis-shaped and the nuclei are small. The non- degenerated L1 layer is marked with a star. (E) The ovule arrests at FG1 stage, while the overall ovule shape reaches later stages. (F) The ovule arrests at FG2 stage, while the overall ovule shape reaches later stages. The non- degenerated L1 layer is marked with a star. (F’) Enlarged view of embryo sac from H. (G) Ovule arrests at FG3 stage, while the overall ovule shape reaches later stages. Ovule stages are determined according to Schneitz et al., 1995.

35S:NTAP:AtCTF7∆B leads to megaspore alterations and degeneration Other types of degrading megaspores and defective embryo sacs were also observed (Figure 2.13). Some ovules contained degrading or completely degraded megaspore(s) at various stages

(Figure 2.13A-D). Unexpectedly, some megaspores appeared to omit meiosis II (Figure2.13 E-H). In some instances the MMC underwent meiosis I to produce a dyad (Figure 2.13E&F). But the resulting megaspore at the basal end seemed to undergo mitosis instead of meiosis II to produce two or more nuclei in the same cell (Figure 2.13F&H, F’&H’). Some of these defects may be caused by improper MMC division and further investigations are required to verify the identities of the megaspores’ (haploid or diploid) and the properties of the divisions (meiotic or mitotic). The majority of the meiotic defects in ovules occured in meiosis II, which is consistent with the observation that the most severe male meiocyte defects were detected during meiosis II. More defects occurred in the embryo sacs during megagametophyte-associate mitosis, such as degrading nuclei (Figure 2.13J&K), totally degenerated embryo sacs (Figure 2.13M), polar nuclei fusion defects (Figure 2.13N) and vacuole development defects (Figure 2.13O&P). The mitotic defects in the ovules might be directly due to reduced AtCTF7 activity or accumulated defects, which occurred in meiosis.

Figure 2.13 35S:NTAP:AtCTF7∆B ovules contain various types of defective gametophytes during meiosis and mitosis. (A-D) 35S:NTAP:AtCTF7∆B ovules contain degrading/degraded megaspore(s) at various stages, revealed by CLSM. (A) Ovule contains two degrading megaspores at the dyad stage after meiosis I. (B) Ovule contains a DM but the associated megaspore contains no nucleus (arrow), indicating the nucleus is completely degraded. (C) Ovule contains a mis-shaped, degrading nucleus in the FM(L). (D) Ovules do not contain visible megaspore(s), indicating the megaspore(s) are completely degraded. (E-H) Megaspores omit meiosis II, revealed by CLSM (E,F,F’) and DIC (G,H,H’). (E) Two megaspores after meiosis I are elongated in shape and contain a diffuse signal throughout the cells but no defined nuclei. (F) The distal megaspore (arrow) becomes a DM. The megaspore at the chalazal end, marked as a star, contains multiple nuclei inside one cell. The nuclei of the chalazal end megaspore are not separated by a cell wall, so the division is mitotic. (G) Two megaspores after meiosis I are elongated in shape. (H) The basal megaspore contains two nuclei in the same cell (marked as a star), while the distal megaspore persists. F’ and H’ are the enlarged views of the megaspores of F and H. (I-K) 35S:NTAP:AtCTF7∆B ovules contain degenerating nuclei during mitosis, revealed by DIC. (I) Ovule arrests at FG1 stage with defects around the FM. (J) Ovule contains one degenerating nucleus (arrow) and one surviving nucleus. The non- degenerated L1 layer is marked with a star. (K) Ovule contains two degrading nuclei (arrows) on the side of the vacuole, other nuclei are not visible. The ovule shape reaches the FG5 stage. J’ and K’ are the enlarged views of the megaspores of J and K. (L-P) Mature 35S:NTAP:AtCTF7∆B ovules contain defective embryo sacs, revealed by DIC. (L) Mature WT ovule. (M) Ovule contains the debris of an embryo sac (arrow). (N) Embryo sac produces two nuclei, morphologically similar to ENs. No vacuole forms at the

basal end. (O) Embryo sac contains the EN and the CN, while the EN and CN are not separated as in WT ovules. No vacuole forms at the basal end. (P) Embryo sacs contain the EN and the CN and the EN and CN are not separated. A huge vacuole appears at the basal end. AN, antipodal nucleus; CN, central nucleus; DM, degenerated megaspore; EN, egg nucleus; FM(L), functional megaspore (like); MMC, megaspore mother cell; SN, synergid nucleus; V, vacuole. FM(L)s are identified as having distinctly bright autofluorescence in the nuclei and the DMs contain diffuse signal throughout the cells but no clearly defined nucleus (Barrell and Grossniklaus, 2005). Ovule stages in DIC are determined according to Schneitz et al, and in CLSM are defined according to Christensen et al., 1998. Bar scale=10 μm.

Slow female gametophyte development and altered megaspores are common phenotypes To validate the ovule defects are caused by high-level expression of the 35S:NTAP:AtCTF7∆B construct, mutant ovules from other severely reduced fertility lines (# 13 and # 15 ) were examined. Similar to Line 11, the gametophytes from these lines developed slowly (Figure 2.14) and asynchronously, and embryo sac maturation was delayed. The majority of the megaspores from these lines developed similar to Line 11, even though the embryo sacs arrested at different stages (Figure 2.14A,C). Unexpectedly, some ovules contained other types of defects, including the co-existence of two FMLs and ovules where the middle megaspores appear to be FMLs (Figure 2.14B,D). Therefore, overexpression of 35S:NTAP:AtCTF7∆B leads to elongated meiosis and asynchronous development of the ovules, along with defective megaspores/embryo sacs. Moreover, tetrad stage was not observed in 35S:NTAP:AtCTF7∆B ovules, suggesting that meiosis was not complete in the mutant ovules or that the mutant ovules formed various FMLs. For comparison, in wild type ovules 8.5% of the ovules are at dyad stage, 26.3% of the ovules are at triad stage and 65.2% of the ovules are at tetrad stage when ovules are at a similar size.

Figure 2.14 Female gametophytes from other lines develop slowly and some megaspores alter their identities. (A) Slowed gametophyte development in Line 13. (i) Ovule only contains megaspore(s). (ii) The ovule is enlarged and a megaspore separates from the other megaspore(s). (iii) The separated megaspore localizes at the more chalazal position and becomes a FML. (iv) The remaining megaspore(s) at the distal end divides and the resulting megaspores degraded. (B) Two FMLs co-exist in some ovules of Line 13. (i) Ovule contains a megaspore with bright fluorescence. (ii) Dyad after meiosis I containing strong fluorescence throughout the cells. (iii, iii’) Two FMLs co-exist at the chalazal end; both contain brightly stained nuclei. The two FMLs are separated and one contains an enlarged nucleus. (iii’) Enlarged view of the megaspores of iii. (C) Slowed gametophyte development in Line 15. (i) Ovule contains a separated megaspore and clustered megaspore(s). (ii,iii) The separated

chalazal-end megaspore becomes a FML. It appears to complete one round of mitosis to produce two nuclei and a vacuole forms between the two nuclei. (D) The middle megaspores become FMLs in some ovules of Line 15. (i) Ovule only contains a FML (arrow) as its nucleus is brightly stained but no associated DM(s) are identified, revealed by CLSM. (ii-iii) Middle megaspores (arrows) are FMLs while the associated megaspores (stars) are degrading, by DIC. The developmental stages are defined according to Christensen et al. 1998. Bar scale=10 μm.

2.2.4 High level expression of NTAP:AtCTF7∆B disrupts the expression of genes involved in female gametophyte development and other related processes Since female gametophyte development was impaired in 35S:NTAP:AtCTF7∆B plants, quantitative reverse transcription polymerase chain reaction (qRT-PCR) was carried out to examine transcript levels for several genes important for ovule development (Figure 2.15A). Transcript levels of WUS1 (Gross-Hardt et al., 2002), DMC1 (Couteau et al., 1999), SYN3 (Jiang et al., 2007; Yuan et al., 2012), OSD1 (d'Erfurth et al., 2009) and MMD1 (Yang et al., 2003) were all increased in buds of 35S:NTAP:AtCTF7∆B plants The greatest increase was observed for WUS1.

Figure 2.15 Elevated expression levels of genes involved in female gametophytes in 35S:NTAP:AtCTF7∆B plants. Siliques of wild type and non-dwarf 35S:NTAP:AtCTF7∆B Line 11 plants were used. Tublin was used as the control. Transcript levels were measured using RT-qPCR. Data are shown as means ± SD (n = 3).

Transcript levels of cohesin genes in 35S:NTAP:AtCTF7∆B plants were also measured. Even though the distribution of SYN1 was not affected (Figure 2.7), SYN1 transcript levels were decreased to to approximately 30% of wild type (Figure 2.16). SYN2/SYN4 expression levels were significantly increased in 35S:NTAP:AtCTF7∆B plants, while SMC1 and SMC3 transcript levels were unchanged (Figure 2.16B). The altered expression levels of SYN1 and SYN2/SYN4 suggested that cohesion may be affected in 35S:NTAP:AtCTF7∆B plants. The opposite changes in SYN1 and SYN2/SYN4 expression levels suggest that 35S:NTAP:AtCTF7∆B may affect the meiotic and mitotic cohesion differently. Transcript levels of a number of genes involved in DNA segregation and recombination, including Rad51, Spo11-1, ZYP1a and 1b, RCK1, PHS1, MLH1 and MSH4 were also analyzed (Figure 2.16B). While transcript levels of SPO11-1 and ZYP1a were found to be higher in 35S:NTAP:AtCTF7∆B plants, transcript levels of other genes were similar to wild type.

Figure 2.16 Altered expression levels of genes involved in cohesion, synapsis and recombination in 35S:NTAP:AtCTF7∆B plants. (A) Expression analysis of cohesin genes in 35S:NTAP:AtCTF7∆B and wild-type plants. (B) Expression of several genes involved in synapsis and recombination is altered in 35S:NTAP:AtCTF7∆B plants. Siliques of wild type and non-dwarf 35S:NTAP:AtCTF7∆B plants (Line 11) were used. Tublin was used as the control. Transcript levels were measured using RT-qPCR. Data are shown as means ± SD (n = 3).

2.3 Discussion

Previous studies demonstrated that reducing AtCTF7 levels by T-DNA mutations or AtCTF7- RNAi leads to sister chromatid cohesion loss along with other defects in male meiocytes (Bolanos-Villegas et al., 2013; Singh et al., 2013). Comparably, the effects of 35S:NTAP:AtCTF7∆B on male meiocytes are less severe and the defects occur at later stages. In 35S:NTAP:AtCTF7∆B meiocytes, the first noticeable defect is chromosome fragmentation during telophase I. Sister chromatid cohesion loss is observed in meiosis II instead of early meiosis I as in atctf7 plants. The milder defects in 35S:NTAP:AtCTF7∆B male meiocytes is consistent with the presence functional AtCTF7 protein and indicates that AtCTF7∆B protein does not interfere with early stages of sister chromatid cohesion. Native AtCTF7 mRNA levels are about three-fold higher in 35S:NTAP:AtCTF7∆B plants than in WT plants, suggesting that NTAP:AtCTF7∆B may interfere with the function(s) of AtCTF7. It also suggests that plant cells may monitor sister chromatid cohesion and contain a feedback mechanism to regulate AtCTF7 levels. Previous studies demonstrated that AtCTF7 functions critically in ovule development (Jiang et al., 2010; Singh et al., 2013). Even though different phenotypes are observed when AtCTF7 levels vary, each mutation seems to arrest the embryo sac at one stage or very few stages. Conversely, 35S:NTAP:AtCTF7∆B causes pleiotropical ovule/seed defects, suggesting different levels of AtCTF7∆B are present or different pathways are disrupted. In severely reduced fertility lines, overexpression of NTAP:AtCTF7∆B leads to asynchronous ovule development, additional gametic cells, megaspore identity alterations, delayed embryo sac maturation and various defective megaspores/embryo sacs. Even though alterations are commonly observed during pre- meiosis and meiosis, the megaspores can progress beyond FG1, suggesting the defects may recover or megaspore identity can be established at later stages. Furthermore, tetrad stage is not observed in some lines, suggesting that meiosis is not complete and the mutant ovules form various defective FMLs. Recent studies have shown that embryo sacs are enriched in transcripts for genes encoding proteins involved in RNA metabolism and transcriptional regulation, and display distinct epigenetic patterns (Wuest et al., 2010; Schmidt et al., 2011; Shi and Yang, 2011). Disruption of genes in small RNA regulatory pathways, such as ARGONAUTE1 (AGO1), AGO 9, DICER-

LIKE1 (DCL1 and MEIOSIS ARRESTED AT LEPTOTENE1 (MEL1), leads to multiple megaspore-like cells at premeiosis, abnormally divided megaspores during meiosis, gametic cell fate alterations and twin female gametophytes (Nonomura et al., 2007; Olmedo-Monfil et al., 2010; Shi and Yang, 2011). For example, AGO9 participates in small RNA silencing by cleaving endogenous mRNAs (Olmedo-Monfil et al., 2010). Ago9 mutations result in additional gametic cells in pre-meiotic ovules, which may skip meiosis and twin female gametophytes in the post- meiotic ovules. Similar defects are observed in 35S:NTAP:AtCTF7∆B ovules during pre-meiosis and meiosis. Twin female gametophytes were not detected in 35S:NTAP:AtCTF7∆B ovules even though similar defects are observed surronding 1-nuclear (1NFG) female gametophytes. Moreover, AGO9 participates in TE silencing. In 35S:NTAP:AtCTF7∆B plants, transcript levels of TEs are dramatically increased and transcript levels of several genes involved in RNA processing are altered. The similar phenotypes between 35S:NTAP:AtCTF7∆B and Ago9 highly suggests small RNA mediated pathways are disrupted in 35S:NTAP:AtCTF7∆B plants. Compared to 35S:NTAP:AtCTF7∆B plants, Ago9 plants are fertile and Ago9 seeds are viable, indicating other pathways may also be affected in 35S:NTAP:AtCTF7∆B plants beyond the small RNA regulatory pathways. Previous studies showed that mutations in Eco1/Ctf7 result in defects in nucleolar integrity, rRNA production, ribosome biogenesis and protein biosynthesis in Saccharomyces cerevisiae and human (Gard et al., 2009; Bose et al., 2012). In Arabidopsis, mutations in genes participating mRNA maturation, rRNA biogenesis and ribosome biogenesis slow the mitotic progression of female gametophytes and result in embryo sacs with pleiotropic defects in megaspore identities and alterations, and polar nuclear fusion defects (Tzafrir et al., 2004; Shi and Yang, 2011; Shi et al., 2005; Coury et al., 2007; Groß-Hardt et al., 2007; Huang et al., 2010; Schmidt et al., 2011;Szakonyi and Byrne, 2011; Zsögön et al., 2014). For example, SLOWWALKER1 (SWA1,) participates in 18S pre-rRNA processing. A mutation in swa1 leads to complete female sterility and asynchronous megagametophyte development, and results in embryo sacs that arrest over a wide range of stages (Shi et al., 2005). Similar defects are also observed in 35S:NTAP:AtCTF7∆B ovules. Some of the 35S:NTAP:AtCTF7∆B ovule defects are also similar to those observed in atsyn3- ovules. Embryo sacs in Atsyn3-2- ovules develop asynchronously and contain various defects, including variable numbers of nuclei at FG6, nuclear migration defects and delayed senescence (Jiang et al., 2007). Similar defects were

observed in 35S:NTAP:AtCTF7∆B and in atsyn3- ovules (Jiang et al., 2010). SYN1 localization is not affected in 35S:NTAP:AtCTF7∆B meiocytes, similar to the situation in AtSYN3 RNAi meiocytes (Yuan et al., 2012). Similar transcript alterations were also observed for genes involved in the synapsis and recombination. AtSYN3 is an atypical α-kleisin. It mainly localizes in the nucleolus and was predicted to participate in rDNA structure and RNA processing. The similarities among 35S:NTAP:AtCTF7∆B , swa1 and Atsyn3+/- plants suggest that rRNA or ribosome biogenesis might be disrupted in 35S:NTAP:AtCTF7∆B plants. In summary, 35S:NTAP:AtCTF7∆B transgenic plants exhibit multiple alterations in male meiosis, megasporogenesis and megagametogenesis. The different defects may be the result of high level of AtCTF7∆B acting at disruption of different levels, which may include sister chromatid cohesion, epigenetic regulation and rRNA/ribosome biogenesis.

2.4 Materials and methods

Plant materials and growing conditions Arabidopsis thaliana ecotype Columbia was used in this study. To grow plants, seeds were sown in Metro-Mix200 soil (Scotts-Sierra Horticultural Products; http://www.scotts.com), stratified at 4°C in the dark for two days (optional) and put in a growth chamber at 22°C with a 16-h-light/8-h-dark cycle. The transgenic plants were obtained by the floral dip method as described (Clough and Bent 1998). The plants transformed with the bar gene (Phosphinotricin-Acetyltransferase) were selected by spraying with BASTA (glufosinate ammonium) at 200 mg/l after germination. Spraying was repeated three more times with an interval of 4 days. Positive plants were further confirmed by PCR.

Isolation of genomic DNA from plants The fresh leaves from plants were collected before bolting and ground with the plastic pestle in plant extraction buffer (2.5ml of 2M Tris (pH9.5), 500μl of 1M EDTA, 6.25ml of 1M KCl, 40.75ml of sterile water). After centrifugation, the supernatant containing the genomic DNA was mixes with equal amount of isopropanol to precipitate the genomic DNA. The genomic DNA was collected by centrifuge and dried at room temperature. The dried genomic DNA was re- suspended in water or TE for plant genotyping.

Polymerase chain reaction (PCR) techniques PCR was carried out to amplify a PCR product using the plasmid DNA, genomic DNA or complementary DNA (cDNA) as a template. The annealing temperature was based on the melting temperature of the primers (4XG/C)+(2XA/T). Components of PCR were: 10μl 5x PCR buffer, 2μl dNTP Mix (10mM of each dNTP), 2μl Taq DNA polymerase (0.5 unit/ μl), 1μl 10μM Forward Primer, 1μl 10μM Reverse Primer, Xμl RNase-free water to make up 50.0μl total.

Cloning procedures The AtCTF7∆B cDNA fragment (AtCTF7∆299-345, 1-894bp nucleotides,) was removed from the cloned pET22 vector by digestion with NdeI/HindIII and re-cloned into pIADL14 (Jiang et al., 2010). To generate the 35S:NTAP:AtCTF7∆B construct, a 1-894 bp nucleotides fragment was cloned into a Gateway-compatible binary vector containing the NTAP tag (Rohila et al., 2004), a gift from Dr. Qinn Li lab, Miami University. AtCTF7∆B was first PCR-amplified by the primers (1111/1201) and cloned into the pENTR vector. The pENTR vector was then fused with the binary vector by LR recombination reactions (Invitrogen) to make the final 35S:NTAP:AtCTF7∆B construct. The constructs were confirmed by DNA sequencing using the big dye terminator labelling mix method and analyzed by the capillary sequencer ABI3700 (Applied Biosystems). To generate transgenic Arabidopsis plants, each construct was mobilized into Agrobacterium tumefaciens strain AGL-1, and transformed into Arabidopsis thaliana using the floral dip method (Clough and Bent, 1998).

Isolation of total RNA from plants To extract total RNA, buds of either wild type or 35S:NTAP:AtCTF7∆B plants were pooled. For 35S:NTAP:AtCTF7∆B samples, buds were only pooled from the aborted, non-dwarf plants. Fresh plant tissues were frozen in liquid N2 and ground in a pestle and mortar, and then mixed with TRIzol reagent, which contained guanidine thiocyanate and β-mercaptoethanol. Total RNAs were then precipitated by two volumes of 100% ethanol, washed by 70% ethanol and dried at room temperature for 2 hr. The RNA was dissolved in RNase free water and stored at -70 °C.

DNase I treatment of extracted RNA Extracted plant RNA was treated with DNase I at room temperature for 15 minutes to remove genomic DNA before RT-PCR analysis. The mixture was incubated at 65°C for 10 minutes to denature the DNase I. The mixture was extracted by chloroform and centrifuged. The supernatant was transferred to a fresh tube and RNA was precipitated by adding 2 vol 100% ethanol and washed by 70% ethanol and dried at room temperature. RNA was dissolved in RNase free water and stored at -70 °C. DNase I digestion condition: 10X DNase I buffer (Invitrogen) 2μl , RNAsin

(Promega®) 1μl, Turbo DNase I (Ambion)1μl, 10 μg of RNA, RNase Free dH2O xμl, Total mix 20μl

Quantitative reverse transcriptase-polymerase chain reaction (qRT-PCR) RT-PCR was used to measure mRNA levels. For RT-PCR, RNA was first reverse transcribed into complementary DNA (cDNA) by reverse transcriptase with an oligo(dT) primer and a First Strand cDNA Synthesis Kit (Roche, http://www.roche.com). qRT-PCR was performed with the SYBR-Green PCR Mastermix (Bio-Rad, Hercules, CA, USA) and amplification was monitored on a MJR Opticon Continuous Fluorescence Detection System (Bio-Rad). Expression was normalized against β-tubulin-2. The amplification efficiency was assumed to be 1.8 per cycle. At least three biological replicates were performed. The sequences of primers used in these studies are presented in Table 2.5.

Chromosome analysis Whole anthers were dissected from buds and analyzed by staining with Alexander staining (Alexander, 1969). Pollen morphology and viability were compared from flowers of 35S:NTAP:AtCTF7∆B plants, 35S: AtCTF7∆B plants and wild-type plants. For male meiotic chromosome spreads, floral buds were fixed in Carnoy’s fixative

(ethanol:chloroform:acetic acid: 6:3:1). The buds were washed twice in H2O for 15 min and transferred to sodium citrate buffer (10 mM, pH 4.5). The anthers of right sizes were dissected and digested with 1.4% β-glucoromidase, 0.3% cytohelicase, 0.3% pectolyase, 0.3% cellulase in 10 mM sodium citrate for 1 hr in a moist chamber (Ross et al., 1996). The digested anthers were washed with cold 10 mM sodium citrate and cleared in 60% acetic acid. The samples were squashed and frozen on dry ice or in a -70 °C freezer. Before use, the samples were air dried and stained with DAPI (4,6-diamino-2-phenylindole dihydrochloride; 1.5 μg ml−1) (Vector Laboratories, Inc. Burlingame, CA, USA) and then observed with an Olympus 1X81 fluorescence deconvolution microscope system. Images were captured with a Spot camera system (Diagnostic Instruments Inc., http://www.spotimaging.com) and processed. Meiocytes were staged as described previously (Ross et al., 1996).

Immunolocalization studies Floral buds of approximately 0.2-0.6 mm were dissected from the inflorescences under a microscope. Anthers were dissected from the buds, transferred to glass slides and covered with a 0.17mm thick cover slip in water. The meiocytes were released by pressing the cover slip. The released meiocytes were transferred to a poly-lysine coated glass slide and digested in 10 mM sodium citrate (pH 4.5) with 1.4% β-glucoromidase, 0.3% cytohelicase, 0.3% pectolyase, 0.3% cellulase for 30 mins at 37oC in a moist chamber. The digested meiocytes were washed in 1XPBST three times for five mins each. The washed samples were stained with 100μl anti-SYN1 primary antibody (1:500 diluted in blocking solution (1XPBST+ 1% BSA)). The slides were covered with parafilms and incubated overnight at 4 ºC. The next day, the slides were washed in 1XPBST three times for five mins each. The sample was blocked by 100μl secondary antibody (Alexa-488 goat anti-rabbit antibody, 1:1000 diluted in blocking solution) and incubated for 1 hour at room temperature. Finally the slides were washed in 1XPBST five times for five mins each and stained with DAPI. The samples observed with an Olympus 1X81 fluorescence deconvolution microscope system. Images were captured with a Spot camera system (Diagnostic Instruments Inc., http://www.spotimaging.com) and processed.

Ovule analysis of 35S:NTAP:AtCTF7∆B plants and WT plants Inflorescences from 35S:NTAP:AtCTF7∆B plants and WT plants were collected and fixed in 4% glutaraldehyde/12.5 mM cocadylate (pH 6.9) under vacuum for two hrs, followed by overnight fixation at room pressure. The fixed samples were dehydrated in a graded ethanol series (40%, 60%, 80%, 100% stepwisely for one hr each) and cleared in a 2:1 mixture of benzyl benzoate:benzyl alcohol. Ovules from each silique were dissected under a stereo dissecting microscope, mounted and sealed by the coverslips. The ovules were observed on a Zeiss Axioskop microscope under DIC optics as described (Schneitz et al., 1995). Images were collected and processed. The ovules were further viewed by confocal laser scanning microscopy (CSLM) as described (Christensen et al., 1997). The specimens were observed with an argon ion laser (488 nm excitation and 515-530 nm emission) using an Olympus confocal microscope system with a 90i Eclipse microscope and Olympus Flouview 2.0 software (http://www.olympus-global.com/), analyzed with Image Pro Plus (Media Cybernetics; http://www.mediacy.com).

Table 2.5 Primer sequences

Primers Sequence 5’-3’ Purpose Cloning AtCTF7∆B gateway F CACCATGCAAGCCAAAATCAATTC Forward primer for entry clone 35S NTAP CTF7∆B (#1111) AtCTF7∆B gateway R GCTAGTTATTGCTCAGCGG Reverse primer for entry clone 35S NTAP CTF7∆B (#1201)

qRT-PCR To quantify expression Tubulin F TGGATCATGAGTGAGTGAAAAGA Expression control Tubulin R AAAACCACAATGGACAATTTC CTF7 native+O/E F GTT GGG TGA GGA TTG GAT TC CTF7 native+O/E R GCG AGG ATG AGC TCT CTT TT CTF7 native F GTG GGA TTA GAG CGA TTT GG CTF7 native R TTC CTA TGG AGC TTG GTT GTG WUS F TGGATCTATGGAACAAGACTGTT WUS R GGCTTTGCTCTATCGAAGAAGT CLV3 F CGAAGGGTTTAGGACTACATGAAG CLV3 R GTGGGTTCACATGATGGTGCAA SYN3-1 F AAAGAAATTTGGGGCTTTCA SYN3-1 R TGGTGTTCCTACTGGGGAAT SYN3-2 F CGAGTTCGACTTGGAAGATG SYN3-2 R AAGGATCAATGCCAGTAGGAA DMC1 F GTTCATATCAGACCCAAAAAAGCC DMC1 R AGATTCGGAGCATCGTAGACTTTG OSD1 F GAATCTCCGGTGAATCCAGA OSD1R AGAAGGCAACAAACCACCAC MMD1 F TATCCGCGGTATGACTGTGT

MMD1R GCAATAGGGTTCCGATGAAT SYN1 F ATGACATTCCCGAAGGAGTC SYN1 R GCTGGTTGTGGTCTATCCTG SYN2 F ACGTTTTGCCAATCCTTTTC SYN2 R CTGTGGTTGCCCAGTTTTA SYN4 F CTGTGCTTGAACCAGCAACT SYN4 R GACGTATAATGTCGCCATGC SMC3 F ATCCACAGTACCGAACCGCT SMC3 R GTGATGAACTGCGTGCCATAA SMC1 F AGCGATTTCGTGACATGGAAC SMC1 R TGGAGAAGAGCACGCCAA Rad51 F CTTAGGGATGCTGGTCTCTGTAC Rad51 R GTCAACCTTGGCATCACTAATTC Spo11-1 F ATGCATCCATCAGCTTTCAA Spo11-1 R TTTAACCAACCCATCACCAA Spo11-2 F CCACAAGGTTTTTCCTCCAC Spo11-2 R CAAGTCCCATCCCTATGCTT ZYP1a F GAGTGAGGTCTGACAATGAT ZYP1a R CTGCGTTTACATTTTGAGCT ZYP1b F AAGAGCTTTGGTGCAGTTAC ZYP1b R CACTTTGACCGACGAATGTG RCK F GAATGGATGCGAAGTTGTTG RCK R ATTGCCCGTGTAATGTCAGA PHS1 F CATTGTCGACGAATTGAACC PHS1 R CATCTGAGGCCATGTGAAGT MLH1 F AACCCAAGCCACAAAGTCTC MLH1 R CAGCTGTCAACTCCAGCAAT MSH4 F CTCCCTTTCAACAACAGGCA MSH4 R GCGGCAGACTTATTCCTGACA

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Chapter 3: 35S:NTAP:AtCTF7∆B leads to vegetative growth defects

3.0 Abstract In addition to its essential roles in reproduction, AtCTF7 is also important for vegetative growth (Bolaños-Villegas et al., 2013; Singh et al., 2013). Inactivation of Arabidopsis CTF7 (AtCTF7) by T-DNA mutation or a decrease in AtCTF7 mRNA levels leads to severe growth defects. Similarly, patients with RBS show defects including growth retardation, limb reduction and cleft lip/palate or missing fingers/toes (Roberts 1919; Van Den Berg and Francke 1993; Vega et al. 2010). Besides the effects on reproduction, high-level expression of NTAP:AtCTF7∆B caused a wide range of phenotypic alterations in vegetative growth. 35S:NTAP:AtCTF7∆B plants displayed a wide range of vegative defects, including dwarf plants, fused stems, a disruption of phyllotaxis, irregular internodes, floral abnormalities and misshaped leaves. Increased numbers of dwarf plants appeared stochastically in later generations. Furthermore, the transcripts for epigenetically regulated transposable elements (TEs) were elevated in transgenic plants. Taken together, this suggested that high-level expression of AtCTF7∆B disrupted epigenetic regulation. A 35S:AtCTF7∆B construct was made to determine if the defects in 35S:NTAP:AtCTF7∆B plants were caused by AtCTF7∆B or the presence of the N-terminal tag. 35S:AtCTF7∆B plants exhibited reduced fertility and dwarf phenotypes, indicating that the defects in 35S:NTAP:AtCTF7∆B plants were caused by high level expression of AtCTF7∆B. Atctf7-/- plants containing a CTF7promoter:AtCTF7∆B construct were also obtained. The plants grew better than Atctf7 plants, but did not fully complement the Atctf7 phenotype. Therefore, the B motif is required for full AtCTF7 function. In summary, this study further demonstrates that AtCTF7 plays essential roles in vegetative growth and that proper levels of AtCTF7 are critical.

This chapter along with chapter 2 has been submitted to Plant Physiol.

3.1 Introduction and Background In addition to its essential roles in reproduction, AtCTF7 plays important roles in vegetative growth. About 4% of ctf7-/- homozygous seeds germinate; ctf7-/- homozygous plants are slow- growing and are severely dwarf (Bolanos-Villegas et al., 2013). Meanwhile, knockdown of AtCTF7 by RNAi leads to severe inhibition in vegetative growth (Singh et al., 2013). Decreased AtCTF7 levels cause growth defects in aerial and root tissues with the loss of greening. After bolting, decreased AtCTF7 levels leads to inhibition in inflorescence growth and sterility. Similar types of phenotypes are observed in RBS patients, who show growth retardation, limb reduction/asymmetric limb growth, cleft lip/palate or missing fingers/toes and mild-to-severe mental deficiencies (Roberts 1919; Herrmann and Opitz 1977; Van Den Berg and Francke 1993; Vega et al. 2010). ESCO2 appears to function in a dosage-sensitive way as both severe and mildly affected individuals are observed in the same progeny. AtCTF7 functions in vegetative growth have also been tested by analyzing the AtCTF7 promoter using the PAtCTF7nlsGUS reporter (Singh et al., 2013). The AtCTF7 promoter is active in both the shoot and root meristems as well as in young buds and leaves. The AtCTF7 promoter is also active in pollen and the female gametophyte, consistent with the ovule defects that have been observed in Atctf7 mutants. Meanwhile, the AtCTF7 promoter is less active in older leaves and flowers. Therefore, AtCTF7 is required for actively dividing cells in vegetative and reproductive tissues.

3.2 Results

3.2.1 Expression of 35S:NTAP:AtCTF7∆B causes pleiotropic growth defects

During our analysis of the severely reduced fertility 35S:NTAP:AtCTF7∆B lines, we observed that plants displaying vegetative defects appeared in the T2 or T3 generations. Twelve out of the 18 severely reduced fertility lines grew worse in later generations,, including the production of dwarf plants. The observed vegetative abnormalities included dwarf plants, fused stems, disruption of phyllotaxis and irregular internodes (Figure 3.1C-F). The morphological variations between plants of same line were very striking (Figure 3.1). More dwarf plants appeared in later generations. However the dwarf phenotype was not inherited in a Mendelian fashion, with dwarf plants producing both dwarf and non-dwarf progeny (Figure 3.1, Table 3.1). The dwarf plants also varied in morphology and exhibited more severe alternations such as acaulescence, floral abnormalities, homeotic changes and irregular leaves with different architectures and colors (Figure 3.1 and Figure 3.2). Similar to the situation with the dwarf phenotype, the vegetative alterations also appeared stochastically. Plants with mis-shaped leaves also segregated stochastically. Compared to 35S:NTAP:AtCTF7∆B dwarf plants, Atctf7-1 plants were more severely dwarf (Figure 3.2). Most Atctf7-1 plants also exhibited early senescence, which was not observed in the previous study (Bolanos-Villegas et al., 2013) but is consistent with RBS mutations where decreased proliferation capacity in RBS cells is related to cell death (Vega et al., 2010).

Figure 3.1 Progressive morphological aggravation of 35S:NTAP:AtCTF7∆B plants through self- pollination. (A) A wild type Columbia plant. (B) A second generation 35S:NTAP:AtCTF7∆B plant with the severely aborted siliques. The plant has shorter siliques and produces aborted ovules. (C) Third generation 35S:NTAP:AtCTF7∆B plants, containing both dwarf plants and non-dwarf, aborted plants. (D, E) Advanced generation 35S:NTAP:AtCTF7∆B plants (D- fourth generation, E-sixth generation), exhibiting more dwarf plants. Both dwarf and non-dwarf plants exhibit more severe abnormalities. (F) 35S:NTAP:AtCTF7∆B plant with the mildly aborted siliques. The plant is relatively normal, but its siliques contain a reduced number of seeds. The plant is from Line 19. Defects such as reduced apical dominance (arrow), phyllotaxis disturbances (asterisks) and irregular internode length are also denoted. Plant shown in C,D and F. B, C, D, E are progeny for self-pollinated plants of Line 11. Plants shown in C, D, E are progeny of non-dwarf, aborted plants from the previous generation. Also see Table 3.1 for comparison. All plants are approximately 30 days old and are grown under the same environmental conditions. Bar=5 cm.

Figure 3.2 Phenotypes of 35S:NTAP:AtCTF7∆B dwarf plants. The inflorescence defects include acaulescent (B and D), multiple inflorescence branches at the first node (C) and no inflorescence (E and F). The leave defects include aberrant rosette sizes and shapes (A, C, E, F and G), and purple leaves (D and E). Atctf7-1-/- plant in H exhibits an early senescent phenotype. All plants were grown under the same environmental conditions. Plants in A, B, C are 28-days old. Plants in D, E, F, G and H are 40-days old. Scale bar=2 cm.

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Table 3.1 Phenotype variations in 35S:NTAP:AtCTF7∆B plants (Line 11) in different generations.

Non-dwarf (Aborted) Generation Dwarf plants (#,%) plants 2nd plants 100% 3rd plants 24 (18.5%) 106 (81.5%)

4th plants Seeds from dwarf plants 16 (42.1% ) 22 (57.9%) Seeds from non-dwarf plants 25 (45.5%) 30 (54.5%)

5th plants Seeds from dwarf plants 55 (78.6%) 15 (21.4%) Seeds from non-dwarf plants 81 (67.5%) 39 (32.5%)

6th plants Seeds from dwarf plants 55 (81.4%) 13 (18.6 %) Seeds from non-dwarf plants 66 (82.5%) 14 (17.5 %)

35S:NTAP:AtCTF7∆B plants are from Line 11. Seeds from aborted, non-dwarf plants and dwarf plants were collected and sown separately.

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3.2.2 Epigenetic alterations are present in 35S:NTAP:AtCTF7∆B plants

Analysis of epigenetic changes in 35S:NTAP:AtCTF7∆B plants by qRT-PCR The stochastic appearance of the dwarf phenotype and the overall variation in phenotypes suggested that the phenotypes could be the result of epigenetic changes. In order to investigate this possibility qRT-PCR was carried out to measure the expression levels of several epigenetically regulated transposable elements (TEs), including MU1, COPIA28 and soloLTR, and several genes associated with epigenetic events (Moissiard et al., 2012). Expression levels of MU1, COPIA28 and soloLTR were all dramatically increased in 35S:NTAP:AtCTF7∆B plants compared to wild type (Figure 3.3). However, only modest changes were observed in transcript levels of several genes in siRNA processes or chromatin remodeling. Transcript levels of HDA19 and RDM4 were decreased approximately 50% (Figure 3.4A). However, the expression levels of the canonical DNA methylation genes, MET1 and DMT7, (Law et al., 2010; Law et al., 2011; Ausin et al., 2009, Chen et al., 2010; Henderson and Jacobsen, 2008) did not vary much. The expression levels of TEs and related epigenetic genes did not vary significantly between dwarf and non-dwarf plants (data not shown). Also, no significant expression differences were detected from different generations of the same line (data not shown). Altogether, our data suggest that AtCTF7 may participate in epigenetic regulation. Transcript levels of cell cycle genes, RBR, CYCB1.1 and CYCA1.1, and the DNA repair genes, BRCA1 and BRCA2B, were also tested and found to be elevated (Figure 3.4B). The similar expression patterns and morphological defects observed between 35S:NTAP:AtCTF7∆B plants and Atctf7-1 plants (Bolanos-Villegas et al., 2013) suggest that similar pathways are being affected.

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Figure 3.3 The mRNA levels of the epigenetically regulated transposable elements, MU1, COPIA28 and soloLTR, are dramatically increased in 35S:NTAP:AtCTF7∆B plants. Siliques of wild type and non-dwarf 35S:NTAP:AtCTF7∆B plants were used. Tublin was used as the control. Data are shown as means ± SD (n = 3) and are from three biological samples.

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Figure 3.4 Transcriptional analysis of genes involved in epigenetics, cell cycle and DNA repair. Figure 3.4A Several genes participating in siRNA process exhibit disturbed expression. Transcription levels of HDA19 and RDM4 are decreased, while MET1 and DMT7 transcription levels are not altered. Figure 3.4B The expression levels of cell cycle genes, CYCB1.1, CYCA1.1 and RBR, and DNA repair genes, BRCA1 and BRCA2B, are significantly increased. Siliques of wild and non-dwarf 35S:NTAP:AtCTF7∆B plants were used. Data are shown as means ± SD (n = 3) and are from three biological samples.

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In general, our study suggests that AtCTF7 is involved in meristem maintenance and differentiation. AtCTF7 appears to act in a dosage-sensitive way since different transgenic lines exhibit varying degrees of defects. 35S:NTAP:AtCTF7∆B appears to lead to epigenetic changes as the morphological defects vary and the dwarf phenotype appears stochastically, which is consistent with qRT-PCR results. An accumulation of defects were also observed in Atctf7+/- plants as fewer heterozygous Atctf7+/- plants are obtained in later generations of self-pollinated Atctf7+/- plants (Jiang et al., 2010).

DNA methylation does not removed from retroelements To analyze the effects of 35S:NTAP:AtCTF7∆B on the methylation status of the genome, the methylation status of the well-characterized retroelements, AtSN1 and IgN5, were tested. Genomic DNA from wild type, 35S:NTAP:AtCTF7∆B, 35S:AtCTF7∆B (addressed in later section) and Atctf7 plants were isolated and digested by the methylation-sensitive enzymes HaeIII and MspI followed by PCR (Figure 3.5). The results showed that IgN5 was not completely de-methylated or completely methylated in 35S:NTAP:AtCTF7∆B, 35S:AtCTF7∆B and Atctf7 plants as they were not cleaved by HaeIII or MspI. Consistent with the results above, cytosine methylation at CG, CHG, and CHH sites in IgN25, IgN5 and IgN23 elements were not altered consistently in Atctf7 (Bolaños-Villegas unpublished data). Therefore, the epigenetic alterations in 35S:NTAP:AtCTF7∆B plants do not appear to be caused by DNA methylation changes, which is consistent with qRT-PCR results showing that expression levels of the canonical DNA methylation genes, MET1 and DMT7, did not vary significantly.

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Figure 3.5 DNA methylation is not completely removed from IgN5 transposons. Genomic DNA from wild type, 35S:NTAP:AtCTF7∆B, 35S:AtCTF7∆B and Atctf7 plants were subjected to PCR (#0), or digested with the methylation-sensitive enzyme HaeIII (#1) or MspI (#2) followed by PCR. Note: HaeIII digests GGCC but not GG mCC; MspI digests GG mCC but not GGCC.

3.2.3 Analysis of 35S:AtCTF7∆B and CTF7:AtCTF∆B plants Our study shows that high level expression of the NTAP:AtCTF7∆B construct has a strong effect on the plant fertility and growth. We next wanted to determine if these effects are due to the presence of the N-terminal tag or the presence of high levels of protein missing the acetyletransferase B motif. Therefore, a 35S:AtCTF7∆B construct was generated (Figure 3.6) and transformed into WT Col plants. Eight out of 13 35S:AtCTF7∆B transgenic plants examined exhibited reduced fertility, with the 35S:AtCTF7∆B defects appearing more severe than those in 35S:NTAP:AtCTF7∆B plants. For example, the dwarf plants occurred at a higher frequency and the phenotypes of the dwarf plants varied more significantly (Figure 3.7B, Line 21). Furthermore, fourth generation seeds from Line 21 were not viable. Similar abnormalities were observed between 35S:NTAP:AtCTF7∆B and 35S:AtCTF7∆B plants, along with additional alterations in 35S:AtCTF7∆B plants. For example, 35S:AtCTF7∆B Line 24 segregated for two types of plants: plants without an inflorescence and reduced fertile plants (Figure 3.7C). 35S:AtCTF7∆B Line 29 plants displayed a unique phenotype in that the siliques were downward-pointing instead of normal upward-pointing (Figure 3.7D). The siliques of Line 29 plants contained fewer seeds (46.4±2.5 versus 54.2±4.1 per silique in WT; n =35) but the seeds germinated normally.

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Therefore, the defects observed in 35S:NTAP:AtCTF7∆B plants appear to be caused by high level expression of AtCTF7∆B and not the NTAP tag. In fact the presence of the NTAP appears to reduce the severity of the alterations, possibly by reducing the stability of the protein or affecting its activity. To determine if the B motif is essential for AtCTF7 function, a CTF7pro:AtCTF7∆B construct was created (Figure 3.6) and transformed into Atctf7-1+/- plants. Wild type plants transformed with CTF7pro:AtCTF7∆B did not display vegetative defects or severe reduced fertility defects (52.0±2.2, n=33 versus 54.2±4.1 per silique in WT; n =35). Atctf7-1-/- plants containing the CTF7pro:AtCTF7∆B construct were obtained at low frequencies (6/86). The plants were generally dwarf but grew better than Atctf7-1-/- plants (Figure 3.8A). Their morphology also varied with acaulescent or fewer rosette leaves defects present in the lines. The siliques contained aborted ovules resembling to Atctf7-1-/- plants, but the plants produced approximately 40 ovules per silique (Figure 3.8C). In summary, the B motif is required for full AtCTF7 function as the CTF7pro:AtCTF7∆B construct only partially restored Atctf7-1-/- defects in vegetative growth and in ovule development.

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Figure 3.6 AtCTF7 constructs used in this study. The AtCTF7∆B construct missing the acetyltransferase B motif and expressed under 35S promoter and AtCTF7 promoter; AtCTF7 (1-177aa) and AtCTF7 (143-345aa) constructs missing the C terminus and N terminus respectively; AtCTF7(C98Y C101Y) construct contains mutations in the C2H2 zinc finger motif.

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Figure 3.7 Morphological variations of 35S:AtCTF7∆B plants. (A) Wild type plant. (B) 35S:AtCTF7∆B plants from Line 21. The dwarf plants appear at a high frequency with various severity. Note: Second generation plants are reduced fertile but not dwarf. (C) 35S:AtCTF7∆B plants of Line 24. The same pot produces reduced fertile plants and plants without an inflorescence. The plants without an inflorescence produce more leaves and the leaves are in normal shape. (D) 35S:AtCTF7∆B plant of Line 29. The plant contains multiple inflorescence branches from the ground and produces downward-pointing siliques which contain a reduced number of seeds (46.4 ±2.5 versus 54.2±4.1 per silique in WT plants; n =35). All plants are 30-day old and grown under the same environmental conditions. 35S:AtCTF7∆B plants are third generation. Scale bar= 5cm.

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Figure 3.8 Phenotypes of Atctf7-1-/- plants containing the CTF7pro:AtCTF7∆B construct. (A) Homozygous Atctf7-1-/- plants containing the CTF7pro:AtCTF7∆B cconstruct. The plants are dwarf. Some plants (1) are stemless and some plants (3) produce less rosettes. (B) Wild type silique with full seed set. (C) A silique from Atctf7-1-/- plant containing the CTF7pro:AtCTF7∆B construct. The silique contains about 40 aborted ovules. All plants are 30-days old and grown under the same environmental conditions. The 35S:AtCTF7∆B plants are 3rd generation. Scale bar= 5cm in A. Scale bar=0.5 cm in B/C.

Other constructs, including 35S:AtCTF7 N-term (1-177aa), 35S:AtCTF7 C-term (143-345aa) and 35S:AtCTF7 C98Y C101Y (Figure 3.6) were created and transformed into WT Col plants. The constructs did not cause reduced fertility or detectable growth defects. Therefore no further investigation was carried out on the lines.

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3.3 Discussion

35S:NTAP:AtCTF7∆B plants exhibit various growth defects related to meristem disruption, which is consistent with the previous result that AtCTF7 is highly expressed in meristematic areas (Singh et al., 2013). The plants also exhibited apparent epigenetic alterations such as the stochastic appearance of traits and the loss of transcriptional silencing of TEs. Morphological variations between plants from the same pot were very striking. However morphological variations are also present in Atctf7-/- plants (Bolaños-Villegas et al., 2013). The milder defects observed in 35S:NTAP:AtCTF7∆B plants provide better opportunities to fully explore the functions of AtCTF7. The apparent epigenetic effect of 35S:NTAP:AtCTF7∆B was demonstrated by the dramatically increased transcript corresponding to normally silenced TEs. Therefore, our study is the first time to demonstrate that Eco1/Ctf7 participates in the epigenetic control, which will certainly expand Eco1/Ctf7 function in gene regulation. The plants transformed with the 35S:AtCTF7∆B construct also exhibited reduced fertility and dwarf phenotypes, indicating that the defects in 35S:NTAP:AtCTF7∆B plants were caused by high level expression of AtCTF7∆B and not the presence of the NTAP on the protein. Among the 35S:AtCTF7∆B lines, Line 29 produced downward-pointing siliques, which was similar to bp/knat1 mutations (Shi et al., 2011; Sakakibara et al., 2013), suggesting that AtCTF7 may be involvled in the KNOX2 pathway. Deletion of KNOX2 genes in the moss Physcomitrella patens leads to gametophyte development without meiosis (Sakakibara et al., 2013), which is consistent with the meiosis II omission phenotype in some 35S:NTAP:AtCTF7∆B ovules. Atctf7-1-/- plants transformed with CTF7pro:AtCTF7∆B were dwarf and ovule development was partially recovered, suggesting that the motif B is essential for AtCTF7 function.

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3.4 Materials and methods

Plant materials and growing conditions The wild type Arabidopsis thaliana (ecotype Columbia) plants and T-DNA insertion line, SALK_059500 (ctf7-1), were chosen in this study. T-DNA insertion plants were genotyped by PCR using primer pairs specific for the T-DNA and wild-type loci. Seeds were snown on Metro- Mix200 soil (Scotts-Sierra Horticultural Products; http://www.scotts.com), stratified at 4°C in the dark for two days (optional) and put in a growth chamber at 22°C with a 16-h-light/8-h-dark cycle. The transgenic plants were obtained by the floral dip method as described (Clough and Bent 1998). The plants transformed with the bar gene (Phosphinotricin-Acetyltransferase) were selected by spraying with BASTA (glufosinate ammonium) at 200 mg/l after germination. Spraying was repeated three more times with an interval of 4 days. The positive plants were further confirmed by PCR.

Isolation of genomic DNA from plants The fresh leaves were collected before bolting and ground with the plastic pestle in plant extraction buffer. After centrifugation, the supernatant containing the genomic DNA was mixed with an equal amount of isopropanol to precipitate the genomic DNA. The genomic DNA was collected by centrifugation and dried at room temperature. The dried genomic DNA was re- suspended in water or TE for plant genotyping or Chop-PCR.

Polymerase chain reaction (PCR) techniques PCR was carried out to amplify a PCR product using either plasmid DNA, genomic DNA or complementary DNA (cDNA) as a template. The annealing temperature was based on the melting temperature of the primers (4XG/C)+(2XA/T).

Chop-PCR for DNA methylation assays For Chop-PCR was carried out as previously described (He et al., 2009). Extracted genomic DNA (500 ng) was digested by HaeIII or MspI, methylation-sensitive enzymes, overnight at

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37°C. After digestion, about 10% of the DNA was used as the template to test methylation patterns at the AtSN1 and IgN5 loci using gene-specific primers. PCR conditions were 4 min at 94°C followed by 35 cycles of 30 sec at 94°C, 30 sec at 56°C, and 1 min at 72°C. PCR products were then subjected to electrophoresis in a 1.2% agrose gel.

Cloning procedures for the constructs To clone 35S:AtCTF7∆B, a 1-894bp fragment containing AtCTF7 sequence except the last B motif, was amplified with primers 1010/1427. The PCR product was digested by NcoI/SpeI and ligated into binary vector pFGC5941. The AtCTF7 promoter, comprising 1.5kb of DNA upstream of the ATG, was amplified with primers 1256/1257, digested by EcoRI/NcoI and cloned into EcoRI/NcoI digested 35S:AtCTF7∆B construct. 35S:N-term (1-177aa) AtCTF7 and 35S:C-term (143-345aa) AtCTF7 was amplified with primers 1244/1245 and 1246/1247, respectively, and ligated into binary vector pFGC5941. 35S:AtCTF7 C98Y C101Y was generated using QuikChange® Site-Directed Mutagenesis Kit with primers 1258/1259. Restriction enzymes were obtained from New England Biolabs. Plasmid DNA and PCR amplified fragments were digested at 37oC between 1 and 4 hours in appropriate buffers, depending on the amount of DNA and the amount of enzymes. The DNA products were analyzed by gel electrophoresis and isolated. All constructs were confirmed by DNA sequencing using the big dye terminator labelling mix method and analyzed by the capillary sequencer ABI3700 (Applied Biosystems). To generate the transgenic Arabidopsis plants, each construct was mobilized into Agrobacterium tumefaciens strain AGL-1, and transformed into Arabidopsis thaliana using the floral dip method (Clough and Bent, 1998).

Isolation of total RNA from plants To extract total RNA, buds of WT and 35S:NTAP:AtCTF7∆B plants were pooled. For 35S:NTAP:AtCTF7∆B samples, buds were only pooled from reduced fertile, non-dwarf plants.

Fresh plant tissues were frozen in liquid N2 and ground in a pestle and mortar, and then mixed with TRIzol reagent, which contained guanidine thiocyanate and β-mercaptoethanol. Total RNAs were then precipitated by 2 vol of 100% ethanol, washed by 70% ethanol and dried at room temperature for 2 hr. The RNA was dissolved in RNase free water and stored at -70 °C.

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DNase I treatment of extracted RNA Extracted plant RNA was treated with DNase I to remove genomic DNA before RT-PCR analysis. The DNase I digestion mixture was kept at room temperature for 15 minutes. Then the mixture was incubated at 65°C for 10 minutes to denature the DNase I. The heated mixture was extracted by chloroform and centrifuged. The supernatant was transferred to a fresh tube and RNA was precipitated by adding 2 vol 100% ethanol and washed by 70% ethanol and dried at room temperature. RNA was dissolved in RNase free water and stored at -70 °C.

Quantitative reverse transcriptase-polymerase chain reaction (qRT-PCR) RT-PCR was used to detect expression of mRNA. For RT-PCR, RNA was first reverse transcribed into complementary DNA (cDNA) by reverse transcriptase with an oligo(dT) primer and a First Strand cDNA Synthesis Kit (Roche, http://www.roche.com). qRT-PCR was performed with SYBR-Green PCR Mastermix (Bio-Rad, Hercules, CA, USA) and amplification was monitored on a MJR Opticon Continuous Fluorescence Detection System (Bio-Rad). Expression was normalized against β-Tubulin-2. The amplification efficiency was assumed to be 1.8 per cycle. At least three biological replicates were performed. The sequences of primers used in these studies are presented in Table 3.2.

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Table 3.2: Primer sequences

Primers Sequence 5’-3’ Purpose Genotyping SALK059500LP ATGAATAGAAATGAGCGATAGGC Genomic forward SALK059500RP GAAGCTATGAGCTTAAATGCTTCC Genomic reverse SALK LBb1.3 ATTTTGCCGATTTCGGAAC Left border forward for SALK lines

Cloning AtCTF7 ∆B F GGAGCCATGGATATGC Forward primer for 35S AAGCCAAAATC CTF7∆B (#1010)

AtCTF7 ∆B R GAGACTAGTTTAATTCTAG Reverse primer for 35S TTATTGCTCAGCGG CTF7∆B (#1427)

CTF7pro F CCGGAATTCCACATCCT Forward primer for CTF7pro GGAAATATCTTTGCAA CTF7∆B (#1256)

CTF7pro R GGAGCCATGGCGTCTAGAG Reverse primer for CTF7pro AGAGCTCGAATCCTTGTT CTF7∆B (#1257)

AtCTF7 N- term GGAGCCATGGATATG Forward primer for CAAGCCAAAATC 35S: N- term (1-177aa) (#1244) AtCTF7 N- term GAGACTAGT TAAAT Reverse primer for CCTCACCCAACTC 35S: N- term (1-177aa)

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(#1245)

AtCTF7 C- term F GGATTAATTAAAATGCAAGCCA Forward primer for AAATCAATTCTTTCTTCAAG 35S: C- term (143-345aa) (#1246) AtCTF7 C- term R GAGACTAGTTTAAGAAAAG Reverse primer for TGAGTATCAATTAGCTGA 35S: C- term (143-345aa) (#1247) AtCTF7 C98Y C101Y F CTTCTC AGACATTACGCA Forward primer for GAATATGGAGCTAAGTAT 35S: CTF7 CY mutant (#1258) AtCTF7 C98Y C101Y R ATACTTAGCTCATATTCTGC Reverse primer for GTAATGTCTGAGAAG 35S: CTF7 CY mutant (#1259)

AtSN1 Forward ACCAACGTGCTGTTGGCC CAGTGGTAAATC Chop-PCR AtSN1 Reversed AAAATAAGTGGTGGTTGT ACAAGC

IgN5 Forward TCCCGAGAAGAGTAGAAC AAATGCTAAAA Chop- PCR IgN5 Reversed CTGAGGTATTCCATAGCC CCTGATCC

qRT-PCR To quantify expression Tubulin F TGGATCATGAGTGAGTGAAAAGA Expression control Tubulin R AAAACCACAATGGACAATTTC MU F TAATTTGGCTGACGGAATCAC

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MU R ATTTGGGGGAAAACAAATGAG COPIA28 F AGTCCTTTTGGTTGCTGAACA COPIA28 R CCGGATGTAGCAACATTCACT SoloLTR F AACTAACGTCATTACATACACATCTTG SoloLTR R AATTAGGATCTTGTTTGCCAGCTA HDA19 F GACTGTGATTACAACACACCGT HDA19 R AATTGCCGCCAGTATCCAT AGO1 F TGGACCACCGCAGAGACAAT AGO1 R CATCATACGCTGGAAGACGACT AGO4 F CACTCGCTCTCCTATGTGTACCAAAG AGO4 R CATGGCTTGATGATGTCTCAGACTGATC RDR2 F TGGCGAGAGATAACCGGAGGTATG RDR2 R CTTCTCATCGCGATGGTTTGGATTG DCL3 F GCCTACTTTCGATACCTCGGAAGA DCL3 R GCATACATCACAGCCTCACGATTG NRPD1A F GACTTGTGAAGATGGTTCTGCAGTTG NRPD1A R GTCTTCGAATGTCCCGTCTATTCTTAC MIR 156 F CTCTCCCTCCCTCTCTTTGATTC MIR 156 R AGGCCAAAGAGATCAGCACCGG MIR 172 F TTTCTCAAGCTTTAGGTATTTGTAG MIR 172R TCGGCGGATCCATGGAAGAAAGCTC MET1 F GTGATTCTTAGGGCTATAATGG MET1 R CATTGATGAAGTCCACTTGAC DMT7 F CCCACCTGAGTTTGTGGACT DMT7 R CATTCTGGCCACCATCTCTT RDM4 F ATGGATGGGGTGGGTGAAAG RDM4 R FTAGCACCTTCTTCGGTTTCAC RBR F GGTGGAGGAGAAACTTGTGC RBR R GTGGTTGCTTCCGGTAGTTG CYCA1.1 F GTTTCGGCTGTTGTTTCGAT CYCA1.1 R CATGAGGTCGCTTCTTAGCC

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CYCB1.1 F CCGGAACTGAATCTGCTTAGGA CYCB1.1 R GCTTGGTTCTTCAGCTTCTTCG BRCA1 F TGCTAGAGCCAGAGCTGCAAG BRCA1 R TGGCAACCTGCAATAGAATCC BRCA2B F TCACCTTAAAACCCGCAGTG BRCA2B R CATGGATGACGGATTTGGAA

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3.5 References Ausin I, Mockler TC, Chory J, Jacobsen SE (2009) IDN1 and IDN2 are required for de novo DNA methylation in Arabidopsis thaliana. Nat Struct Mol Biol 16: 1325101038/nsmb1690 Bolanos-Villegas P, Yang X, Wang H, Juan C, Chuang M, Makaroff CA, Jauh G (2013) Arabidopsis CHROMOSOME TRANSMISSION FIDELITY 7 (AtCTF7/ECO1) is required for DNA repair, mitosis and meiosis. Plant J 75: 927–940 Chen PY, Cokus SJ, Pellegrini M (2010) BS Seeker, precise mapping for bisulfite sequencing. BMC Bioinformatics 11: 203101186/1471-2105-11-203 Clough SJ, Bent AF (1998) Floral dip, a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J 16(6): 735–743 He X, Hsu Y, Pontes O, Zhu J, Lu J, Bressan RA, Pikaard C, Wang C, Zhu J (2009) NRPD4, a protein related to the RPB4 subunit of RNA polymerase II, is a component of RNA polymerases IV and V and is required for RNA-directed DNA methylation. Genes Dev 23: 318-330 Heidinger-Pauli JM, Unal E, Guacci V, Koshland D (2008) The kleisin subunit of cohesin dictates damage-induced cohesion. Mol Cell 31: 47–56 Jiang L, Yuan L, Xia M, Makaroff CA (2010) Proper levels of the Arabidopsis cohesion establishment factor CTF7 are essential for embryo and megagametophyte, but not endosperm, development. Plant Physiol 154(2): 820-32 Law JA, Jacobsen SE (2010) Establishing, maintaining and modifying DNA methylation patterns in plants and animals. Nat Rev Genet 11: 204101038/nrg2719 Law JA, Vashisht AA, Wohlschlegel JA, Jacobsen SE (2011) SHH1, a homeodomain protein required for DNA methylation, as well as RDR2, RDM4, and chromatin remodeling factors, associate with RNA polymerase IV. PLoS Genet 7: e1002195101371 Moldovan GL, Pfander B, Jentsch S (2006) PCNA controls establishment of sister chromatid cohesion during S-phase. Mol Cell 23: 723-732 Roberts J (1919) A child with double cleft of lip and palate, protrusion of the intermaxillary portion of the upper jaw and imperfect development of the bones of the four extremeties. Ann Surg 70: 252–253 Sakakibara K, Ando S, Yip HK, Tamada Y, Hiwatashi Y, Murata T, Deguchi H, Hasebe M,

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Bowman JL (2013) KNOX2 genes regulate the haploid-to-diploid morphological transition in land plants. Science 339(6123):1067-70 Shi C, Stenvik GE, Butenko MA (2011) Arabidopsis Class I KNOTTED-Like homeobox proteins act downstream in the IDA-HAE/HSL2 floral abscission signaling pathway. Plant Cell 23: 2553–2567 Singh DK, Andreuzza S, Panoli AP, Siddiqi I (2013) AtCTF7 is required for establishment of sister chromatid cohesion and association of cohesin with chromatin during meiosis in Arabidopsis. BMC Plant Biology 13:117-124 Van Den Berg DJ, Francke U (1993) Roberts Syndrome, a review of 100 cases and a new rating system for severity. Am J Med Genet 47: 1104–1123 Vega H, Trainer AH, Gordillo M, Crosier M, Kayserili H, Skovby F, Uzielli ML, Schnur RE, Jabs EW (2010) Phenotypic variability in 49 cases of ESCO2 mutations, including novel missense and codon deletion in the acetyltransferase domain, correlates with ESCO2 expression and establishes the clinical criteria for Roberts Syndrome. J Med Genet 47(1): 30–7 Vega H, Waisfisz Q, Gordillo M, Sakai N, Yanagihara I, Yamada M, van Gosliga D, Kayserili H, Xu C, Ozono K, Jabs EW, Inui K, Joenje H (2005) Roberts Syndrome is caused by mutations in ESCO2, a human homolog of yeast ECO1 that is essential for the establishment of sister chromatid cohesion. Nat Genet 37: 468–470

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Chapter 4: Conclusions and Perspectives

Conclusions

Eco1/Ctf7 is essential for the establishment of sister chromatid cohesion during S phase of the cell cycle (for reviews, see Nasmyth and Haering, 2009; Brooker and Berkowitz, 2014; Zamariola, et al., 2014). Knockout of Ctf7/Eco1 leads to a lethal phenotype in most organisms. Altering Eco1/Ctf7 levels or point mutations in the gene can lead to significant alterations in nuclear division as well as a wide range of developmental defects (reviewed in Dorsett and

Merkenschlager, 2013). Eco1/Ctf7 contains a C2H2 zinc finger motif in the N-terminal portion of the protein and an acetyltransferase domain in the C-terminal portion (Dyda et al., 2000; Ivanov et al., 2002). Mutations in the N-terminus of the gene typically lead to severe defects in cohesion and often to chromosome loss during mitosis (Brands and Skibbens, 2005). Meanwhile certain mutations in the C-terminal acetyltransferase domain have less of an effect on cohesion and chromosome segregation, but often result in an increased sensitivity to DNA-damaging agents (Lu et al., 2010). Similar phenotypes are observed in Arabidopsis thaliana when AtCTF7 levels are varied. Heterozygous AtCtf7+/- plants grow normally but approximately 25% of the seeds are aborted, consistent with the notion that CTF7 is an essential protein in plants (Jiang et al., 2010). Unexpectedly, homozygous atctf7 plants are detected at very low frequencies (Bolanos-Villegas et al., 2013). atctf7 plants exhibit extreme dwarfism and sterility along with a wide range of developmental defects. Knockdown of AtCTF7 mRNA levels also causes defects in sister chromatid cohesion and growth retardation (Singh et al., 2013).

In this thesis, it was shown that high-level expression of NTAP:AtCTF7∆299-345 results in reduced fertility and severe defects in vegetative growth. Specifically, 35S: NTAP: AtCTF7∆B plants exhibit defects in male and female meiocytes. Female reproduction is most dramatically affected. Male meiocytes exhibited chromosome fragmentation and uneven chromosome segregation during meiosis II, which ultimately resulted in pollen abortion. Various defects were observed in ovules, including abnormal megasporocyte-like cells at pre-meiosis, megaspores that experienced elongated and aborted meiosis, and defective megaspores and embryo sacs at various stages. A wide range of vegetative defects in inflorescence development, leaf and plant stature were also observed in AtCTF7∆B transgenic plants. These defects appeared stochastically and were not inherited in a Mendelian fashion.

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Comparison of reproductive defects in 35S:NTAP:AtCTF7∆B, AtCTF7 RNAi and atctf7 plants AtCTF7 RNAi and atctf7 plants display severe defects in chromosome condensation and sister chromatid cohesion during early meiotic prophase along with defects in homologous chromosome pairing and segregation later in meiosis (Bolanos-Villegas et al., 2013; Singh et al., 2013). Comparably, male meiocytes from 35S:NTAP:AtCTF7∆B plants were less affected and the majority of defects occured in meiosis II as twenty or more individual chromosomes were often observed starting at metaphase II, suggesting cohesion was prematurely lost. Overexpression of AtCTF7∆B did not disturb the distribution of SYN1 on meiotic chromosomes suggesting that the initial establishment of cohesion was normal. Likewise, no noticeable defects were observed in chromosome condensation, sister chromatid cohesion and homologous chromosome pairing during meiotic prophase in 35S:NTAP:AtCTF7∆B plants. This suggests that AtCTF7∆B does not affect the bulk of meiotic cohesin complexes to a significant extent, but rather may alter centromeric cohesin levels, or possibly cohesin interactions with SGO1 or PATRONUS (Cromer et al., 2013). In contrast, the effect(s) of 35S:NTAP:AtCTF7∆B on female reproduction are observed earlier and are more variable than those in atctf7 and AtCTF7 RNAi plants. 35S:AtCTF7 or AtCTF7 RNAi blocks early ovule development, typically at FG1 or FG2 (Jiang et al., 2010; Singh et al., 2013). In contrast, atctf7 ovules from AtCTF7+/- plants develop relatively normally, but arrest soon after fertilization (Jiang et al., 2010). In all three situations the ovules arrest at a specific developmental stage with relatively uniform alterations in each case. In contrast, 35S:NTAP:AtCTF7∆B causes pleiotropic ovule/seed defects. High level expression of NTAP:AtCTF7∆B led to a wide range of defects, including abnormal cells adjacent to the gametic cells, delayed/arrested meiosis, the production of functional megaspore-like cells of which some were mis-positioned, and various defective embryo sacs, along with defective seeds. Although alterations were commonly first observed prior to and during meiosis, most megaspores progressed beyond FG2 before arresting. It is not clear if the alterartions observed in early stages and those in late stages are related and caused by the same mechanism(s). Previous studies have shown that mutations in Eco1/Ctf7 lead to defects in nucleolar integrity, rRNA production, ribosome biogenesis and protein biosynthesis in Saccharomyces

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cerevisiae and human (Gard et al., 2009; Bose et al., 2012). In Arabidopsis, mutations in genes participating in mRNA production and rRNA/ribosome biogenesis slow mitotic progression in female gametophytes and result in pleiotropic defects in embryo sacs (Tzafrir et al., 2004; Shi et al., 2005; Coury et al., 2007; Groß-Hardt et al., 2007; Huang et al., 2010; Schmidt et al., 2011; Shi and Yang, 2011; Szakonyi and Byrne, 2011; Zsögön et al., 2014). For example, mutations in SLOW-WALKER1 (SWA1), which participates in 18S pre-rRNA processing, result in asynchronous megagametophyte development, and embryo sacs arrest over a wide range of stages (Shi et al., 2005). Defects in 35S:NTAP:AtCTF7∆B ovules are similar to those of Atsyn3- ovules. Embryo sacs in Atsyn3-2- ovules develop asynchronously with defects in the numbers of nuclei, position of nuclei and nuclei senescence (Jiang et al., 2007). AtSYN3 was postulated to participate in rDNA structure and RNA processing (Jiang et al., 2007). Likewise, mutations in ribosomal protein genes leads to defects in inflorescence, leaf and plant development in Arabidopsis similar to those observed in 35S:NTAP:AtCTF7∆B plants (Van Lijsebettens et al., 1994; Byrne, 2002; Stirnberg et al., 2012; Zsögön et al., 2014). Therefore, many of the alterations in 35S:NTAP:AtCTF7∆B plants may be caused by alterations in rRNA or ribosome biogenesis.

35S:NTAP:AtCTF7∆B Leads to Defects Related to Epigenetic Alterations 35S:NTAP:AtCTF7∆B lines displayed relatively normal vegetative growth for the first two generations. However, severe vegetative abnormalities appeared starting in the T2 or T3 generations. The defects, including dwarf plants, fused stems and disrupted phyllotaxis, varied between lines and even between progeny of the same line. The proportion of plants exhibiting vegetative alterations as well as the severity of the vegetative alterations increased in successive generations. Phenotypic variability is consistently observed in RBS patients (Vega et al. 2005, Vega et al., 2010). The increased severity of the vegetative defects and their higher frequency in subsequent generations could result from the accumulation of defects in 35S:NTAP:AtCTF7∆B plants. While the vegetative defects in 35S:NTAP:AtCTF7 plants may in fact result from spontaneous mutations, the situation is clearly more complex as the dwarf phenotype is inherited in a non- Mendelian fashion. When dwarf plants were selfed they produced a mixture of dwarf and non- dwarf, reduced fertile plants. The frequency of dwarf plants produced from selfed dwarf plants

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was similar to the frequency of dwarf plants resulting from selfing a non-dwarf plant. This raised the possibility that the epigenetic state is altered in 35S:NTAP:AtCTF7∆B plants. Consistent with this possibility, transcript levels of MU1, COPIA 28 and solo LTR were increased between five (MU1) and 24 fold (COPIA 28) in 35S:NTAP:AtCTF7∆B plants. Transcript level changes were also observed in several siRNA associated genes, AGO1, AGO4, RDR2, mir156, HDA19 and RDM4. The possible epigenetic alterations in 35S:NTAP:AtCTF7∆B plants are consistent with the alterations in ovule development. Recent studies have shown that the embryo sac displays distinct epigenetic regulatory mechanisms, and is enriched for transcripts involved in RNA metabolism and transcriptional regulation (Wuest et al., 2010; Schmidt et al., 2011; Shi and Yang, 2011). Disruption of genes in small RNA regulatory pathways, such as ARGONAUTE1 (AGO1), AGO9, DICER-LIKE1 (DCL1) and MEIOSIS ARRESTED AT LEPTOTENE1 (MEL1), leads to multiple gametic cells at premeiosis, abnormal meiotic divisions, gametic cell fate alterations and twin female gametophytes (Nonomura et al., 2007; Olmedo-Monfil et al., 2010; Shi and Yang, 2011). For example, mutations in AGO9, which participates in small RNA silencing by cleaving endogenous mRNAs, result in additional gametic cells in pre-meiotic ovules, which may skip meiosis, and twin female gametophytes in post-meiotic ovules (Olmedo-Monfil et al., 2010). Similar defects are also observed in 35S:NTAP:AtCTF7∆B ovules during pre-meiosis and meiosis. Moreover, AGO9 participates in the epigenetically regulated silencing of TEs (Olmedo- Monfil et al., 2010), which were also elevated in 35S:NTAP:AtCTF7∆B plants. The epigenetic alterations in 35S:NTAP:AtCTF7∆B plants may be directly related to NTAP:AtCTF7∆B overexpression or a secondary effect. In the first case, NTAP:AtCTF7∆B may directly affect the expression of genes involved in epigenetic regulation. In the later case, the epigenetic alterations may be an indirect effect. For example, cohesin complexes mediate the transcriptional regulation of many genes, some of which are involved in epigenetic regulation and can have far-ranging developmental consequences (reviewed in Dorsett and Merkenschlager, 2013). Therefore, over expression of NTAP:AtCTF7∆B may modifiy cohesin complexes or other factors to cause the epigenetic alterations.

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Perspectives: AtCTF7∆B likely acts on several levels

In 35S:NTAP:AtCTF7∆B plants, the transcript levels of NTAP:AtCTF7∆B and native AtCTF7 are both elevated, indicating the 35S:AtCTF7∆B construct does not cause co-suppression. Therefore, high-level expression of AtCTF7∆B appears to exert a dominant negative effect. Less clear is how high level expression of AtCTF7∆B exerts its effect or if all of the alterations are related. There are a couple of ways AtCTF7∆B may act. High-level expression of AtCTF7∆B may directly compete with native AtCTF7 for substrates or for interacting factors. To obtain a general idea of what aspects are affected, transcriptional profiling experiments could be performed to test the expression levels of genes between 35S:NTAP:AtCTF7∆B plants and wild type plants. Eco1/Ctf7 acetylates a number of substrates including Eco1/Ctf7 itself, Smc3, Mcd1/Scc1, Irr1/Scc3, Pds5 and Msp3 (reviewed in Skibbens, 2009; Xiong and Gerton, 2010; Ghosh et al., 2012). Deletion of the last 46 amino acids of the acetyltransferase domain is expected to dramatically alter its actyltransferase activity, and therefore alter the acetylation states of various substrates. Previous studies showed cohesin complexes are regulated by acetylation. The differentially acetylated cohesin complexes will affect the interactions between the cohesin complexes and chromosomes during loading, cohesion establishment and cohesion removal. Further experiments such as CHIP (chromatin immunoprecipitation) could be performed to compare the binding patterns of cohesin complexes on chromosomes in 35S:NTAP:AtCTF7∆B and in wild type plants. In addition to its roles in sister chromatid cohesion, cohesin complexes also fuction in other processes including transcription and translation (for reviews, see Nasmyth and Haering, 2009; Yuan et al., 2011; Bose T et al., 2012; Brooker and Berkowitz, 2014; Zamariola, et al., 2014). During transcriptional regulation, cohesin can fold or tether distant DNA chromosomal loci together by interacting with transcription factors, including CCCTC-binding factors (CTCFs) and mediators (Sofueva and Hadjur, 2012). Techniques such as FISH (Fluorescence in situ hybridization) and ChIA-PET (Chromatin Interaction Analysis by Paired-End Tag Sequencing) (Fullwood and Ruan, 2009) can be utilized to to study the organization of chromosomes in 35S:NTAP:AtCTF7∆B and wild type plants. In addition to the cohesin complex, Eco1/Ctf7 acetylates other substrates (Skibbens, 2009; Xiong and Gerton, 2010; Ghosh et al., 2012). Previous studies showed that these proteins are

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acetylated differentially during various processes to accomplish their functions. Mutations that h are defective in acetylation cause similar defects as mutations in Eco1/Ctf7 (Ghosh et al., 2012). Therefore some defects observed in 35S:NTAP:AtCTF7∆B plants may be caused by the altered acetylation of non-cohesin-complex substrates. Further experiments, such as MALDI (Matrix- assisted laser desorption/ionization) mass spectrometry, can be carried out to analyze the acetylation patterns of the different substrates in different plants. Finally, the possibility also exists that the C-terminal deletion may alter the interaction of ECO1/CTF7 with other proteins, either directly or indirectly involved in maintaining chromatin structure. ECO1/CTF7 interacts with various factors, including RFCs (components of the clamp loader replication factor C) (Kenna and Skibbens, 2003), CTF4 and CTF18 (Lengronne et al., 2004), and PCNA (DNA polymerase processivity factor) (Moldovan et al., 2006). Yeast two hybrid and coimmunoprecipitation (Co-IP) could be performed to study the interactions between AtCTF7∆B/ AtCTF7 and other factors. Further experiments are required to determine how specifically AtCTF7∆B is acting, why male and female reproduction respond differently to alterations in AtCTF7 levels and what role, if any AtCTF7 plays in epigenetic regulation. Further analysis of AtCTF7∆B transgenic plants and their comparison with atctf7 and AtCTF7 RNAi plants and CTF7/Eco1 mutants in other organisms will provide interesting insights into potential roles of CTF7.

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