ISOLATION OF COPY NUMBER SUPPRESSORS OF THE NIMA1 KINASE AND MITOTIC REGULATION OF NUCLEOLAR STRUCTURE IN ASPERGILLUS NIDULANS

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree Doctor of Philosophy in the Graduate

School of The Ohio State University

By

Leena Ukil

*****

The Ohio State University 2007

Dissertation Committee: Approved by Dr. Stephen A. Osmani, Advisor

Dr. Berl R. Oakley ______Dr. Hay-Oak Park Advisor Graduate Program in Molecular Genetics Dr. Harold A. Fisk

ABSTRACT

Regulation of the cell cycle is critical for normal development of multicellular organisms and an understanding of this process is crucial for studying cell proliferation and cancer. A number of cell cycle dependent protein kinases specifically control mitotic progression and chromosome segregation. The nimA gene in Aspergillus nidulans encodes one such protein kinase that is both required and sufficient for chromosome condensation, mitotic spindle formation and disassembly of the nuclear pore complex to allow tubulin and regulators to enter nuclei during mitosis. There exist protein kinases structurally similar to nimA in other organisms, including humans. In the filamentous fungus Aspergillus nidulans, the NIMA kinase is required for the regulation of mitosis along with CDK1/cyclin B. Levels of NIMA are regulated throughout the cell cycle, reaching a maximum at mitotic entry and falling dramatically at mitotic exit. Forced expression of the nimA gene can promote mitotic entry, even in human cells. The essential function of NIMA in A. nidulans and the growing recognition of its function in other eukaryotes, means that a study of NIMA function would reveal unique insights into cell cycle regulation among a broad range of organisms. I describe here the characterization of three novel genes mcnA, mcnB and mcnC, three multi-copy number suppressors of the nimA1 conditional mutant, identified in a copy number suppression screen of the nimA1 mutant, and describe the potential novel roles they may play in mitotic regulation. Characterization of MCNC suggests that it is involved in mitotic regulation. First, over expressed mcnC suppresses the G2 arrest caused by nimA1. Second, MCNC over expression leads to the dispersion of the nuclear pore protein SONB suggesting its potential role in regulating NIMA localization to the nucleus during mitosis. Third, mcnC genetically interacts with nimA. Additionally, the deletion of mcnC causes polarization defects due to delayed germtube emergence. Over expression of MCNC concomitantly

ii leads to multiple germ tube formation suggesting a positive regulatory role of mcnA in controlling polarized growth of A. nidulans. The other two nimA1 suppressing genes, mcnB and mcnA both lead to up regulation of NIMA protein levels when over expressed. mcnB is nuclear during G2/M and carries a prominent transcription factor domain called the forkhead domain. MCNB begins to accumulate in the nucleus during G2 and peaks at mitotic entry suggesting it may play a role as a nimA specific transcription factor. Its ortholog in Schizosaccharomyces pombe, Sep1, is known to specifically regulate transcription of a number of genes with roles in sister chromatid separation, septation and cytokinesis and shown to be required for the periodic accumulation of the nimA related kinase fin1. Therefore, a role for forkhead transcription factors regulating cell cycle specific nimA expression is conserved between S. pombe and A. nidulans. MCNA was found to have a fascinating and dynamic location within cells. Endogenously tagged mcnA appears as a single dot in the nucleus. Co-localization studies of MCNA with nucleolar markers show MCNA to locate in the vicinity of the nucleolus and to have a unique pattern of segregation during mitosis. During G2, MCNA is closely associated with nucleolar proteins and appears as an intranuclear body. During mitotic DNA segregation, the MCNA body localizes to the cytoplasm. When mitosis is completed, MCNA remains as a single body outside newly formed daughter nuclei which begins to appear within new daughter nuclei only during G1. At the same time, multiple smaller MCNA bodies are formed in the cytoplasm which finally disappears to completely re-accumulate into the two daughter nuclei by very late G1. Our studies of MCNA localization through the cell cycle shows a highly specific pattern for MCNA and puts forth suggests a possible function of mcnA in regulating nimA turnover during mitosis. The association of MCNA with the nucleolus prompted us to further study the nucleolus in A. nidulans and its segregation during mitosis. The nucleolus is a prominent nuclear structure whose mitotic segregation is poorly understood. During yeast mitosis the nucleolus segregates intact with rDNA (the nucleolar organizing region – NOR). In contrast, during open mitosis the nucleolus is disassembled then reassembled during

iii mitosis. In A. nidulans nuclei, mitosis is a partially open process and I demonstrate that the nucleolus segregates through a completely novel mechanism. Unlike Saccharomyces cerevisiae, few A. nidulans nucleolar proteins segregate with DNA. Instead during DNA segregation, a double pinch of the NE occurs that results in the formation of two daughter nuclei and a central tertiary cytoplasmic structure we have termed the nuclear remnant that contains several nucleolar proteins. While the NOR segregates with the rest of the DNA, the bulk of the nucleolar proteins are seen to remain distinctly intact in the cytoplasm within this nuclear remnant structure. It is only during late telophase and early G1 that the nucleolar proteins from the remnant structure begin to undergo a sequential disassembly and reassembly into the daughter nuclei resulting in the formation of two functional daughter nucleoli in a step-wise manner. My study indicates that nucleolar disassembly-reassembly in A. nidulans is under the control of the spindle assembly checkpoint and that the step-wise process may be regulated by the preferential action of the mitotic kinase CDK1 towards certain nucleolar proteins. A potential role of the phosphatase BIMG is also suggested as BIMG is seen to be localized to the nuclear remnant structure during this process. This study also indicates that A. nidulans undergoes mitotic disassembly then reassembly of its nucleolus, as do higher eukaryotes, and that generation of daughter nuclei occurs via a double fission mechanism, not a single fission as occurs in yeasts. I suggest this novel mitotic nuclear remnant serves as a storage pool from which equal distribution of nucleolar proteins can occur. Mathematical modeling provides supportive evidence for this hypothesis. The nuclear remnant may also serve as a sink for unwanted cytoplasmic proteins or RNAs that gain access to the nucleoli during partially open mitosis.

iv

DEDICATION

This work is dedicated to my parents,

to my brother Shubha,

and to my loving husband Bose.

v

ACKNOWLEDGMENTS

I am deeply indebted to my advisor Dr. Steve Osmani for his continual guidance, training and support throughout the years of my graduate studies. This achievement would not have been possible without his extreme patience and motivation.

I would also like to thank my committee members, Dr. Berl Oakley, Dr. Hay-Oak Park, Dr. Harold Fisk and Dr. Russell Hill for their time and guidance during my graduate studies at The Ohio State University.

Lastly, I would like to thank all current and former members of the Osmani lab for their friendship and support thus making my experience in the lab enjoyable. I would also like to specially thank Aysha Osmani for her motherly care and guidance and for making me a part of her extended family.

vi

VITA

1997-2001……………………………….B.S. Biochemistry, University of Madras.

2002 – 2003……………………………..Teaching Asst. The Ohio State University.

2003 – 2007……………………………..Research Asst. The Ohio State University.

PUBLICATIONS

1. Nayak, T., Szewczyk, E., Oakley, C.E., Osmani, A., Ukil, L., Murray, S.L., Hynes, M.J., Osmani, S.A., Oakley, B.R. (2006). A versatile and efficient gene-targeting system for Aspergillus nidulans. Genetics 172 : 1557- 66.

2. Yang, L., Ukil, L., Osmani, A., Nahm, F., Davies, J., De Souza, C.P., Dou, X., Perez-Balaguer. A., Osmani, S.A. (2004). Rapid production of gene replacement constructs and generation of a green fluorescent protein-tagged centromeric marker in Aspergillus nidulans. Eukaryotic Cell. 3 : 1359-62.

3. Dou, X., Wu, D., An, W., Davies, J., Hashmi, S.B., Ukil, L., Osmani, S.A. (2003). The PHOA and PHOB cyclin-dependent kinases perform an essential function in Aspergillus nidulans. Genetics 165: 1105-15.

FIELDS OF STUDY

Major Field: Molecular Genetics

vii

TABLE OF CONTENTS

Page ABSTRACT……………………………………………………………………………..ii DEDICATION…………………………………………………………………………v ACKNOWLEDGMENTS………………………………………………………………vi VITA……………………………………………………………….. …………………vii LIST OF FIGURES………………………………...... …………………xiv LIST OF TABLES………………………………………………………………...…xvii

CHAPTERS:

1. INTRODUCTION...... 1

1.1. Project Goal...... 1 1.2 Aspergillus nidulans...... 2 1.2.1. General classification and description...... 2 1.2.2. Life cycle of Aspergillus nidulans...... 3 1.2.2.1. Asexual reproduction...... 3 1.2.2.2 Sexual reproduction...... 3 1.2.2.3 The parasexual cycle...... 4 1.2.3. A.nidulans as a model organism for cell cycle research...... 5 1.3. The Cell Cycle...... 7 1.3.1. Introduction...... 7 1.3.2 Interphase...... 8 1.3.3. Mitosis phase...... 9 1.3.3.1. The three flavors of mitosis...... 10 1.3.4. Regulation of the cell cycle...... 11 1.3.4.1. Key cell cycle discoveries...... 11 1.3.4.2. Checkpoint controls exist to monitor proper cell cycle progression...... 11 1.3.4.3. Cyclin dependent kinases (CDK)...... 13 1.3.4.3.1. Mitotic regulation by CDK1...... 13 1.4. NIMA protein kinase...... 14 1.4.1 Functional domains and important residues in NIMA...... 15 1.4.2. NIMA levels through the cell cycle...... 17 1.4.3. NIMA protein kinase is hyperphosphorylated during mitosis...... 18 1.4.4. NIMA interacting proteins and targets...... 19 1.4.5. NIMA related kinases and their role in the cell cycle...... 20 viii 1.5. The Nucleolus...... 23 1.5.1. Repeated copies of the ribosomal RNA genes are the organizers of the nucleolus...... 24 1.5.2. Chromosomal location of repeated copies of ribosomal genes...... 26 1.5.3. Ribosomal gene transcription is required for formation of the nucleolus formation...... 26 1.5.4. Molecular architecture of the Nucleolus...... 28 1.5.5. Nucleolar constituents are highly dynamic...... 29 1.5.6. Nucleolar proteomic studies: new information regarding nucleolar content...... 30 1.5.7. Maintenance of nucleolar structure can be uncoupled from ribosomal RNA transcription...... 31 1.5.8. Nucleolar structure during mitosis...... 32

2. MATERIAL AND METHODS...... 43

2.1. General DNA preparation and cloning...... 43 2.1.1. Plasmid maxiprep and miniprep...... 43 2.1.2. DNA cloning...... 43 2.1.3. Polymerase chain reaction (PCR)...... 44 2.1.4. Primers...... 44 2.1.5. DNA sequencing ...... 44 2.1.6. Bacterial strains...... 45 2.1.7. Transformation of bacteria...... 45 2.1.8. Storage and stock preparation of bacteria...... 45 2.2. Culture and genetics of A. nidulans...... 46 2.2.1. A. nidulans specific media...... 46 2.2.2. Preparation of A. nidulans conidia stock suspensions...... 47 2.2.3. Conidiospore Quantitation...... 47 2.2.4. Long term Storage and Stock Preparation of A. nidulans...... 48 2.2.5. Strain generation by meiotic crossing...... 48 2.3. General A. nidulans techniques...... 49 2.3.1. Small scale protein preparation...... 49 2.3.2. Large scale protein preparation...... 49 2.3.3. Small scale genomic DNA extraction...... 50 2.3.4. Large scale genomic DNA extraction...... 51 2.3.5. Transformation of A. nidulans ...... 52 2.3.6. Immunofluorescence...... 53 2.4. Isolation of full-length cDNAs by 5’RACE PCR...... 54 2.5. Fusion PCR...... 55 2.5.1. Gene deletion constructs...... 55 2.5.2. Endogenous C-terminal tagging constructs...... 56 2.6. S-tag purification in A. nidulans...... 57 2.7. Coomassie staining of protein gels...... 58

ix 2.8. Silver staining of protein gels...... 58 2.9. In vitro λ-phosphatase assay...... 59 2.10. alcA driven protein expression in A. nidulans...... 59 2.11. Measuring NIMA1 levels in nimA1 suppressed strains...... 60 2.12. Western blot analysis...... 61 2.13. Site-directed mutagenesis...... 61 2.14. ΔmcnA, ΔmcnB and ΔmcnC phenotype testing...... 62 2.15. Crosses between ΔmcnC and cell cycle mutants...... 62 2.16. Examination of polarization defects in ΔmcnC expressing cells...... 63 2.17. Microscopy and image capture software...... 63 2.18. Bioinformatics and DNA analysis...... 63

3. COPY NUMBER SUPPRESSORS OF THE nimA1 MUTANT...... 72

3.1. Introduction...... 72 3.1.1. Conditional mutants...... 73 3.1.2. Genetic screens and identification of conditional ts mutants...... 74 3.1.3. The nimA ts mutants...... 76 3.1.4. High copy number suppression of nimA1 – the AMA1 system...... 78 3.1.4.1. The utility of suppressor screens...... 78 3.1.4.2. The AMA1 plasmid...... 80 3.2. Results...... 81 3.2.1. Isolation of copy number suppressors – screen and experimental design...... 81 3.2.2. Isolation of suppressor plasmids from suppressors...... 82 3.2.3. Confirmation of non-nimA suppressors by transformation in the nimA5 strain...... 84 3.2.4. Sequence characterization and sub-cloning of nimA1 suppressor...... 84 3.2.5. The multi-copy number suppressor of nimA1 genes mcnA, mcnB and mcnC...... 85 3.2.6. The effect of over-expression of mcn genes on NIMA protein profile...... 86 3.3. Discussion...... 88 3.3.1. nimA1 suppression by over expression of NIMA1 protein levels..88 3.3.2. nimA1 suppression by modification of NIMA1 protein localization...... 89

4. MCNC GENETICALLY INTERACTS WITH NIMA...... 98

4.1. Introduction...... 98 4.1.1. NIMA interacting proteins...... 98 4.2. Results...... 99 4.2.1. mcnC sequence characterization...... 99 4.2.2. MCNC protein is non-uniformly localized to the cytoplasm...... 100

x 4.2.3. MCNA protein runs during SDS PAGE at a higher size than predicted...... 100 4.2.4. mcnC function is required for normal growth...... 101 4.2.5. Characterization of the ΔmcnC growth defects...... 101 4.2.6. mcnC over expression affects polarity of the cells...... 102 4.2.7. MCNC over expression affects nuclear pore protein localization103 4.2.8 mcnC genetically interacts with nimA...... 103 4.3 Discussion...... 104 4.3.1. mcnC over expression affects the nuclear pore complex protein SONB...... 104 4.3.2. mcnC genetically interacts with nimA...... 105 4.3.3. Positive regulation of cell growth by mcnC...... 105

5. MCNB IS A FORK-HEAD DOMAIN CONTAINING PUTATIVE TRANSCRIPTION FACTOR...... 117

5.1. Introduction...... 117 5.1.1. nimA specific transcription factors...... 117 5.1.2. Fork-head domain containing transcription factors...... 118 5.2. Results...... 120 5.2.1. Sequence characterization of mcnB reveals a forkhead domain...120 5.2.2. MCNB localizes to the nucleus during G2-M transition...... 120 5.2.3. mcnB is a non-essential gene...... 121 5.3. Discussion...... 121 5.3.1. mcnB up-regulation of NIMA1 is likely due to increased transcription...... 121 5.3.2. MCNB and NIMA show identical protein expression profiles....122 5.3.3. Functional redundancy of forkhead transcription factors...... 122

6. MCNA IS A NOVEL SUB-NUCLEAR PHOSPHOPROTEIN WHICH TRANSIENTLY LOCATES TO CYTOPLASMIC BODIES SPECIFICALLY DURING MITOSIS AND G1 OF THE CELL CYCLE...... 128

6.1. Introduction...... 128 6.1.1. Regulation of gene function through protein/mRNA turnover...... 128 6.1.2. Regulation of protein activity by nucleolar sequestration...... 129 6.1.3. Programmed degradation of mRNA take place in P bodies...... 129 6.1.4. Cyclin B mRNA is degraded during mitosis via a specialized P body...... 130 6.2. Results...... 131 6.2.1. mcnA sequence reveals a novel protein specific to the Aspergilli...... 131 6.2.2. MCNA sub-cellular localization...... 132 6.2.2.1. MCNA localizes to a single focus in the nucleus...... 132 6.2.2.2. The MCNA body is located near the nuclear envelope....132

xi 6.2.3. MCNA location varies through mitosis and G1 of the cell cycle...... 133 6.2.4. MCNA co-localizes near nucleolar proteins in interphase but for only part of mitosis...... 134 6.2.5. S-tag affinity purification of MCNA...... 134 6.2.6. MCNA is a phosphoprotein...... 135 6.2.7. Three potential CDK1 phosphorylation sites of MCNA identified by Mass spectrometric analysis...... 136 6.2.8. mcnA deletion and mutation of MCNA phosphorylation sites...... 136 6.3. Discussion...... 139 6.3.1. MCNA protein localization is cell cycle regulated...... 139 6.3.2 MCNA bodies have some similar features to TAM bodies...... 139 6.3.3 MCNA is cell cycle regulated by phosphorylation...... 140 6.3.4. MCNA over expression leads to NIMA up-regulation: a contradiction?...... 140

7. DURING MITOSIS THE NUCLEOLUS OF ASPERGILLUS NIDULANS UNDERGOES STEPWISE DISASSEMBLY IN THE CYTOPLASM AND IS REASSEMBLED IN G1 NUCLEI...... 148

7.1. Introduction...... 148 7.2. Results...... 153 7.2.1. Mitosis in A. nidulans involves a double pinch of the nuclear envelope resulting in the formation of a transient “nuclear remnant” devoid of DNA...... 153 7.2.2. The nuclear remnant is devoid of DNA...... 154 7.2.3.The nucleolus of A.nidulans segregates during mitosis via a mechanism that generates three “nucleolar” structures...... 155 7.2.4 The transient nuclear remnant formed by the double NE pinch contains the nucleolus but not the nucleolar organizing region (NOR)...... 156 7.2.5. Daughter nuclei re-establish transport before reassembling nucleolar proteins...... 158 7.2.6. Fibrillarin reassembles in daughter nucleoli earlier than Bop1...... 159 7.2.7. Nucleolar disassembly-reassembly is under the control of the spindle assembly checkpoint (SAC)...... 160 7.2.8. Non-degradable cyclin B prevents Bop1 disassembly...... 162 7.2.9. The potential role of BIMG in nucleolar disassembly...... 163 7.2.10. The unique nucleolar segregation pattern of A.nidulans helps to ensure equal distribution of nucleolar proteins to daughter nuclei in a common cytoplasm...... 164 7.2.11. Mathematical modeling of cytoplasmic dispersed protein reassembly into daughter nuclei in a common cytoplasm...... 165

xii 7.2.12. Mathematical modeling of nuclear remnant protein reassembly...... 166 7.3. Discussion...... 167 7.3.1. The mitotic dynamics of the NE in A. nidulans...... 168 7.3.2. The transient nuclear remnant defined by the modified NE contains the nucleolus...... 169 7.3.3. Regulation of disassembly and reassembly of the nucleolus...... 170 7.3.4. Why divide the A. nidulans nucleolus via a cytoplasmic intermediate as opposed to its segregation on the DNA?...... 172

8. FINAL DISCUSSION………………………………………………………………191

8.1. Overview…………………………………………………………………...191 8.2. mcnB probably transcriptionally activates nimA…………………………..192 8.3. mcnC might regulate NIMA sub-cellular localization……………………..193 8.4. mcnA is a novel nucleolar associated protein...... 194 8.5. A new mechanism for mitotic segregation of the nucleolus.………………195

BIBLIOGRAPHY……………………………………………………………………..198

xiii

LIST OF FIGURES

Figure Page

Figure 1.1. Asexual development of A. nidulans...... 33

Figure 1.2. Lifecycles of A. nidulans...... 34

Figure 1.3. Spore color mutants of A. nidulans...... 35

Figure 1.4. Polarized growth and nuclear division in A. nidulans...... 36

Figure 1.5. The eukaryotic cell cycle...... 37

Figure 1.6. nim mutants...... 38

Figure 1.7. NIMA kinase...... 39

Figure 1.8. Regulation of NIMA through the cell cycle...... 40

Figure 1.9. The NIMA-related family of kinases...... 41

Figure 2.1 Generation of a gene deletion construct by 3-way fusion PCR………………65

Figure 2.2 Generation of a C-terminal GFP tagging construct by 3-way fusion PCR…..67

Figure 3.1. Plasmid map of vector pRG3-AMA1-NotI…….……………………………90

Figure 3.2. Outline of copy number suppressor screen of nimA1……………………….91

Figure 3.3. Isolation, purification & grouping of copy number suppressor plasmids…...92

Figure 3.4. Confirmation of non-nimA suppressors by transformation in nimA5……….93

Figure 3.5. Sub-cloning of A13………………………………………………………….94

Figure 3.6. Sub-cloning of D15………………………………………………………….95

Figure 3.7. Sub-cloning of C13…………………………………………………………..96

xiv Figure 3.8. Western blot analysis for NIMA1 protein in copy number suppressed nimA1 strains…………………………………………………………………………………….97

Figure 4.1. MCNC protein sequence…………………………………………………...108

Figure 4.2. Protein localization of MCNC…………………………………………...... 109

Figure 4.3. Protein analysis of MCNC using Western blot…………………………….110

Figure 4.4. Characterization of mcnC deletion…………………………………………111

Figure 4.5. Bright field and DAPI images of ΔmcnC with wild type…………………..112

Figure 4.6. Effect of alcA-mcnC over expression……………………………………...113

Figure 4.7. Effect of over expressed mcnC……………………………………………..114

Figure 4.8. Effect of mcnC over expression on nucleopore protein SONB……………115

Figure 4.9. mcnC genetically interacts with nimA……………………………………..116

Figure 5.1. MCNB protein sequence…………………………………………………...125

Figure 5.2. MCNB protein analysis and localization…………………………………..126

Figure 5.3. MCNB localization through the cell cycle…………………………………127

Figure 6.1. Sequence characterization of MCNA protein………………………………142

Figure 6.2. Sub-cellular localization of MCNA………………………………………...143

Figure 6.3. MCNA-GFP localizes in vicinity of the nucleolus…………………………144

Figure 6.4. MCNA through the cell cycle………………………………………………145

Figure 6.5. MCNA is a phosphoprotein………………………………………………...146

Figure 6.6 Characterization of mutant MCNA-3A……………………………………..147

Figure 7.1. Generation of two daughter nuclei during mitosis in A. nidulans occurs by a double pinch of the nuclear membrane which forms a transient nuclear remnant devoid of DNA...... ……………..175

Figure 7.2. The nucleolus of A. nidulans segregates during mitosis via a mechanism that generates three nucleolar structures…………………………...... 176

xv

Figure 7.3. Mitotic segregation of GFP tagged nucleolar proteins CgrA, Bop1 and Nrap……………………………………………………………………………………..177

Figure 7.4. The transient nuclear remnant formed by double NE pinch contains the nucleolus but not the nuclear organizing region (NOR)………………………………..178

Figure 7.5. Daughter nuclei re-establish transport before reassembling nucleolar proteins………………………………………………………………………………….179

Figure 7.6. Fib-GFP with Bop1-chRED shows step-wise disassembly and reassembly………………………………………………………………………………180

Figure 7.7. The final disassembly of Fib-GFP is under the control of SAC……………181

Figure 7.8. The disassembly of Bop1-GFP is under the control of SAC……………….182

Figure 7.9. The role of cyclin B in nucleolar protein disassembly……………………..183

Figure 7.10. BIMG localization and distribution during mitosis……………………….185

Figure 7.11. Comparative reassembly of NLS-DsRed and nucleolar protein Fib……...186

Figure 7.12A. Mathematical modeling of a Ds-Red like protein that disperses in the cytoplasm on onset of mitosis and is reassembled from the cytoplasm………………..187

Figure 7.12B. Mathematical modeling of a protein restricted to the nuclear remnant during mitosis…………………………………………………………………………..188

Figure 7.13. Summarized model for segregation of nucleolus in A. nidulans………….189

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LIST OF TABLES

Table Page

Table 1.1. List of A. nidulans cell cycle mutants...... 42

Table 2.1. List of primers for gene deletion...... 69

Table 2.2. List of primers for gene tagging...... 70

Table 4.1. mcnC homologs………….…………………………...…………...... 107

Table 5.1. mcnB homologs...... 124

Table 6.1. mcnA homologs...... 142

Table 7.1. Effect of non-degradable cyclin B on Bop1 and Fib disassembly………...184

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LIST OF ABBREVIATIONS

CDK cyclin dependent kinase

NIM never-in-mitosis

DAPI 4',6-diamidino-2-phenylindole

AMA autonomously maintained in Aspergillus

SAC spindle assembly checkpoint

M mitosis

S synthesis

NPC nuclear pore complex

Fib fibrillarin

NE nuclear envelope

MPF maturation promoting factor

APC anaphase promoting complex

SMC structural maintenance of chromosomes

NEK nimA related kinases

NOR nucleolar organizing region

FC fibrillar center

GC granular component

DFC dense fibrillar component

xviii UBF upstream binding factor

SL1 selectivity factor 1

Pol polymerase

PNB pre-nucleolar bodies

RE restriction enzyme

PCR polymerase chain reaction mM millimolar ts temperature sensitive

NCBI national center of biotechnology information

ORF open reading frame

AF A. fumigatus

Δ deletion

P bodies processing bodies

xix

CHAPTER 1

INTRODUCTION

1.1. Project Goal The cell division cycle is an essential process through which growing cells divide, maintain and transfer a complete complement of genetic material to daughter cells. Regulation of the cell division cycle is crucial for normal development of multicellular organisms. The cell cycle consists of distinct phases and the activation of each phase is dependent on the proper progression and completion of the previous one. The basic mechanism of cell growth and division is highly conserved from lower eukaryotes like yeast to higher eukaryotes such as humans. The molecular players regulating the main events in the cell cycle are also fundamentally similar in all eukaryotes. The paradigm of cell cycle regulation through the activation and inactivation of cyclin dependent kinases (CDKs) has been firmly established in all eukaryotes. CDK1, the central regulator of cell cycle progression, plays a major role during mitosis along with specific mitotic steps controlled by polo and aurora kinases. It has been established however that a second class of cell cycle regulated protein kinase, namely NIMA also plays an important role. The observation in Aspergillus nidulans that NIMA is required for mitotic entry in spite of CDK1/cyclin B activation proves the function of NIMA to be indispensable for proper cell cycle progression (Oakley and Morris 1983; Osmani, May et al. 1987; Osmani, McGuire et al. 1991). Additionally, expression of NIMA in yeast, frog or human cells (Lu and Hunter 1995) promote mitotic events while many NIMA related kinases identified in other organisms play similar mitotic specific roles in other eukaryotes. This suggests that a NIMA like pathway may

1 be well conserved in other organisms (O'Connell, Krien et al. 2003; O'Regan, Blot et al. 2007). The identification of NIMA related kinases and many NIMA interacting proteins have provided valuable insights into understanding how NIMA regulates the cell cycle. However, a lot remains to be learnt about the functions and regulation of this protein kinase family along with additional novel roles of NIMA. Questions regarding spatial regulation of NIMA, substrates of NIMA and their identification, molecular consequences of substrate phosphorylations, degradation of NIMA and its target substrates, potential NIMA oligomerization, its activation and inactivation still remain to be answered. An effort therefore to search for additional NIMA interacting proteins and NIMA targets will lead to more insights regarding NIMA and its role in the regulation of the cell cycle. Overall, a better understanding of the cell cycle in the model organism will help us have more insights into the cellular roles of similar proteins in higher eukaryotes.

1.2 Aspergillus nidulans 1.2.1. General classification and description Aspergillus nidulans, also called Emericella nidulans, is a filamentous fungus of the phylum Ascomycota. It is one of the most extensively studied organisms in the field of genetics and biochemistry and was first described as Stergimatocystis nidulans by Eidam in 1883. Aspergilli are ubiquitously found in nature and are commonly isolated from soil, plant debris, and indoor air environment. The genus is well defined and includes important species such as Aspergillus fumigatus, Aspergillus flavus, Aspergillus oryzae and Aspergillus niger. Many species of Aspergillus play important roles in the medicinal, industrial, food and scientific areas. As human pathogens, A. fumigatus and A. flavus can cause a range of fungal infectious diseases collectively termed Aspergillosis. Aspergillus flavus and Aspergillus parasiticus produce a toxic and carcinogenic mycotoxin called Aflatoxin. Aspergillus niger is commercially used in the food industry for the production of citric acids and other organic acids, while Aspergillus oryzae is commonly used to convert plant starch into sugars for the production of fermented food items. Over the past several decades, Aspergillus nidulans has become an important

2 model organism for studies in cell biology and for the purpose of this thesis will be the principle experimental system. Pontecorvo and his team laid the foundation for genetic analysis in the early 1950s (Pontecorvo, Roper et al. 1953). Since then considerable progress has been made in establishing it as a powerful biochemical and genetical system, which allows analysis of a broad range of biological phenomena.

1.2.2. Life cycle of Aspergillus nidulans 1.2.2.1. Asexual reproduction Asexual reproduction in Aspergillus nidulans is initiated by the germination of uninucleate haploid spores called conidia (Figure 1.1A). Under favorable conditions, these spores undergo initial isotropic expansion followed by highly polarized growth forming haploid multi-nucleate germlings (Figure 1.1B). After a few rounds of mitotic divisions, the germling is compartmentalized by the formation of a cross wall, termed septum, which results in a sub-apical cell that exits the cell cycle and an apical cell that continues to grow, divide and form branched hyphae distal to the septa (Fiddy and Trinci 1976). Due to various regulatory pathways, hyphae grown in liquid media do not undergo the asexual developmental program to form conidiophores and hence do not form conidia. On solid media however A .nidulans shows colonial hyphal growth that additionally undergoes conidiation. For conidiation to occur during vegetative hyphal growth, specialized foot cells develop within the hyphae (Figure 1.1C). These foot cells then undergo differentiation to form a stalk like structure. A vesicle is formed at the end of the stalk that gives rise to one or more rows of metulae (primary sterigmata), followed by a row of phialides (secondary sterigmata). The distal nucleus in each secondary sterigmatum undergo rounds of mitotic division followed by cytokinesis resulting in the formation of uninucleate spores (Figure 1.1C) (Todd, Davis et al. 2007). Each condiophore bears as many as a hundred chains of conidia and each chain may contain over a hundred spores. The conidia are the structures that give the colony their corresponding color and appearance. The wild type strains form green spores while several different color mutants exist which form white, yellow, fawn or chartreuse spores (Figure 1.3) (Clutterbuck 1972).

3 1.2.2.2 Sexual reproduction A.nidulans is homothallic, encoding two Aspergillus mating types (Galagan, Calvo et al. 2005) within the same cell, which means that it is self-fertile. Out crosses initiate by hyphal fusions between two vegetative hyphae each carrying nuclei of a particular genetic type. The resulting heterokaryon can undergo a septation event resulting in the separation of two nuclei of different genetic type into one septal compartment. The pair of nuclei destined for meiosis then divides in synchrony to form a mass of cells knows as ascogenous hypha. These hyphae are highly branched and each penultimate cell becomes an ascus in which the two haploid nuclei fuse to form a transient zygote that undergoes meiosis immediately. A subsequent mitotic division results in the formation of eight nuclei, each destined to become an ascospore in an ascus. Each ascospore nucleus then undergoes mitosis to form a mature binucleate ascospore. All the ascospores are contained within the sexual fruiting body (cleistothecia) surrounded by hülle cells that are presumed to function as nurse cells (Todd, Davis et al. 2007). Each cleistothecium can contain as many as 104 ascospores arranged in non linear asci, the progeny of a single ascogenous hypha (Figure 1.2). Vegetative heterokaryons do not necessarily have to undergo sexual development. Usually unstable they can be forced to maintain a balanced representation of each nucleus by including complementing auxotrophic mutations in the parental nuclei and growing them in the absence of the corresponding supplements (Roper 1952).

1.2.2.3 The parasexual cycle In addition to the asexual and sexual cycle, a parasexual cycle exists in A. nidulans. The parasexual cycle initiates when haploid nuclei of a heterokaryon fuse forming diploids and divide mitotically. Haploidization of these diploids can be experimentally induced by treatment with microtubule poisons such as benomyl or Nocodazole. This process of haploidization of diploid nuclei in the absence of meiotic cell division is referred to as the parasexual cycle (Pontecorvo and Kafer 1958). Forming a diploid organism is a genetic tool useful in determining whether mutations are dominant or recessive, in mapping gene order and assign new genes to linkage groups.

4 1.2.3. A.nidulans as a model organism for cell cycle research A.nidulans was first identified by Pontecorvo and colleagues in 1945 as a useful organism with attributes ideal for genetic studies (Pontecorvo, Roper et al. 1953). Since then A.nidulans has become a laboratory organism of high value for research and for understanding genetic principles. Research on A.nidulans has contributed immensely to our knowledge and understanding of various biological and developmental processes including, spore development (Clutterbuck 1969; Adams, Boylan et al. 1988; Mirabito, Adams et al. 1989; Timberlake 1990), cell polarity (Momany 2002), cell cycle (Oakley and Morris 1981; Osmani, Engle et al. 1988; Osmani, Pu et al. 1988; Xiang, Zuo et al. 1999; Osmani and Mirabito 2004), DNA repair (Goldman and Kafer 2004), cell metabolism and its regulation (Brambl 2004), cell signaling (Hicks, Yu et al. 1997), secondary metabolite formation (Yu and Keller 2005) and pH control (Arst and Penalva 2003). A.nidulans is a model filamentous fungus that can be grown and maintained by relatively simple techniques. It grows vegetatively and can undergo both sexual and asexual development. The growth rate of A.nidulans is rapid allowing the generation of large amounts of biomass in a relatively short period of time. A.nidulans is able to grow at a variety of temperatures ranging from 15˚C to 44˚C (Doonan 1994). Mycelium of A.nidulans can be obtained by growing spores in liquid media and can be subsequently used for preparation of DNA, RNA and protein extractions. The compact colonial growth on solid media is an added advantage for laboratory purposes as many individual colonies can be grown on a single petri dish while growth properties like colony diameter, degree of conidiation and hyphal density can be measured. The effects on growth rate by changing composition of the solid media or effects of changing growth conditions (temperature, pH, osmolarity) can also be readily assessed. When grown on rich media has a cycle time of about 100 minutes consisting of a 15 minute G1 phase, a 40 minute S phase, a 40 minute G2 phase and a 5 minute M (Bergen and Morris 1983). The A.nidulans genome contains eight linkage groups (n=8) and a detailed genetic map is available (Clutterbuck 1994). Its genome is 30.07 Mb and contains relatively few repetitive elements. It is haploid allowing easy selection of mutants. The parasexual cycle

5 allows easy mapping of a mutation to the linkage groups (McCully and Forbes 1965; Hastie 1970) while heterokaryons and diploids can be used for the study of dominance using complementation analysis (Pontecorvo, Roper et al. 1953). There is also a vast collection of auxotrophic and color mutants in A.nidulans that can be effectively used as markers during genetic analysis. In addition, due to homothallism, sexual reproduction can yield progeny from a self crosses as well as outcrosses. This makes the need for generating mutants in appropriate mating type background unnecessary. Apart from the above characteristic, early studies showed Aspergillus to exhibit many hallmarks of eukaryotic mitosis (Robinow and Caten 1969). In the early 1970s, Ron Morris successfully utilized these features in setting up a temperature sensitive screen for cell cycle mutants (Table 1.1). Based on direct observation of nuclear and spindle morphology, these mutants were classified into three main groups – those defective in mitosis, those defective for septa formation and those defective for nuclear migration. These mutants were further sub-divided based on characterization of their phenotype. The isolation of 26 unique cell cycle mutants from Morris’s lab provided the foundation for further studies of mitotic regulation in A.nidulans (Osmani and Mirabito 2004). Various other techniques have evolved since then to aid research in the field of cell cycle and regulation. Protocols for visualization of cellular contents such as DNA by DAPI (Figure 1.4) and proteins by immunofluorescence are well established. A.nidulans responds well to traditional arrest treatments using drugs like benomyl and hydroxyurea, allowing for easy arrest of cells at specific cell cycle stages and synchronization of cultures (Bergen and Morris 1983). Additionally, vector based expression systems are available for exogenous protein expression. An AlcA promoter based sequence allows for the repression and induction of protein expression by altering the media carbon source (Waring, May et al. 1989). An autonomously replicating plasmid based on the AMA1 sequence allows for high efficiency complementation of recessive mutations and recovery of the complementing DNA (Aleksenko and Clutterbuck 1996; Aleksenko and Clutterbuck 1997). A robust system also exists for efficient gene deletion and tagging using a fusion PCR technique (Nayak, Szewczyk et al. 2006; Szewczyk, Nayak et al.

6 2006) and the lack of non-homologous recombination in an nKuA deleted strain background to result in a dramatically improved frequency of gene targeting (Nayak, Szewczyk et al. 2006). Given that a variety of techniques are available to effectively study the regulation of cell cycle and given that the genome sequence of eight important asexual Aspergilli are published, A.nidulans is an ideal model organism and provides an opportunity to further our understanding of cell cycle regulation and apply it to higher eukaryotes.

1.3. The Cell Cycle 1.3.1. Introduction The cell cycle is an ordered set of events that collectively leads to cell growth and division forming two daughter cells. The cell cycle also ensures that all genetic material essential for survival of individual cells and the whole organism are properly transmitted from one generation to another. In the past several decades there has been a revolution in the comprehension of how the cell cycle functions in different organisms. Experimental results from various eukaryotes like yeast, Xenopus and mammalian cells have provided us a unified set of principles for the normal cellular growth and division process in plants and animals. The scope of this introduction will be limited to a general description of the different phases of the cell cycle followed by a discussion of its regulation which governs proper cell cycle progression including important breakthroughs on the role of the cyclin dependant kinases, NIMA or NimA kinase and checkpoint controls. The eukaryotic cell cycle can be divided into two broad phase: interphase, a period in which cells grow, accumulate nutrition and double their genetic material followed by mitosis – a period when the cell divides itself and splits its genetic material into two distinct halves. The mitotic phase or M phase is relatively brief and consists of DNA segregation followed by cytoplasmic division (cytokinesis). Interphase on the other hand is the longest part of the cell cycle (Figure 1.5).

7 1.3.2. Interphase Interphase is divided into three phases, G1 (first gap), S (synthesis) and G2 (second gap). Cells grow and mature to attain a critical cell size during G1, followed by the synthesis phase (S) in which the DNA is replicated and doubled. Finally, the G2 phase is where the cells prepare for mitosis. The two gap phases (G1 and G2), involve intensive biochemical activity resulting in a doubling of the cell size with a similar increase in critical enzymes, ribosomes, mitochondria, carbohydrates, structural proteins, lipids, and other biomolecules and organelles that are needed as the cell prepares for mitosis. If the cell is unable to prepare for the S phase due to lack of nutrients or

unfavorable environmental conditions, they can enter a hibernating phase called G0 and exit the cell cycle. They only reenter the cycle after the limiting conditions become favorable. Once the cells exit mitosis they undergo cytokinesis, the process which physically separates the cell into two distinct daughter cells. Although very little activity is observable in the cell nucleus by real-time fluorescence microscopy during interphase (Gasser 2002), we know that the step of duplicating DNA is crucial for cell cycle progression. During S phase a cell must replicate all chromosomes a single time and ensure that the chromosomes are undamaged. The presence of damaged DNA results in an inhibition of mitotic entry until all DNA can be repaired (Schultz, Chehab et al. 2000; Nyberg, Michelson et al. 2002; Cuddihy and O'Connell 2003). The cell must also avoid re-duplication of its DNA prior to mitotic entry. To ensure this, the cell follows precise rules wherein each S phase must be followed by M and re-initiation of chromosome duplication is blocked in S phase and G2 (Heichman and Roberts 1994; Nurse 1994). In all eukaryotic cells, DNA replication is regulated by a set of proteins collectively called the replisome that are recruited to sites within DNA known as “origins”. Origins of DNA replication are therefore the sites in the genome where replication is initiated. The recruitment process, termed initiation, involves the Origin Replication Complex (ORC) that marks the origin and the subsequent loading of the helicase Minichromosome Maintenance complex (MCM) that licenses the DNA for replication at G1. To initiate the replication process, termed elongation, and start DNA replication, two protein kinases, CDK2 (cyclin dependent kinase 2) and DDK

8 (Dbf4 dependent kinase) then activate the helicase and allow the loading of the replisome. The replisome in turn contains DNA polymerase which initiates bidirectional replication of DNA from the origins in a semi-conservative fashion. In addition, re- licensing of the origins and re-replication is prevented by blocking MCM re-loading during S, G2 and M phases by CDK2 (Sclafani and Holzen 2007). This block is then reversed at the start of the next cycle’s G1 by the degradation of mitotic cyclins thereby inactivating the mitotic CDK. The S-CDK not only initiates DNA replication to occur in a cell that is competent for DNA synthesis (G1 cell) but also prevents re-replication of DNA synthesis in a cell (G2 cell) that has already completed S phase. S-CDK is instrumental in firing of the replication origin by phosphorylating one of the inhibitory components cdc6 and causing its degradation, thereby allowing DNA replication to start. At the same time, S-CDK also phosphorylates the MCM protein of the replisome after its firing, causing its export to the cytoplasm thereby preventing reassembly of the replisome and DNA synthesis after S phase. Levels of S-CDK remain high through out the M phase and it is only at the end of M phase when all cyclins are degraded that the next cycle of replisome formation can re occur.

1.3.3. Mitosis phase Mitosis is the phase in which a cell divides its duplicated DNA into two identical units. This is generally followed by cytokinesis which divides the cytoplasm and cell membranes along with daughter nuclei into two distinct separated cells. This phase ensures equal distribution of organelles and other cellular components to the daughter cells. The process of mitosis is highly regulated at each step. The mitotic events can be classified into six sequential but distinct phases. Mitosis begins at prophase which initiates the process causing compaction of the genetic material, the chromatin, into highly ordered structures called chromosomes. Since the genetic material was duplicated in the proceeding S phase, each chromosome actually contains a pair of sister chromatids bound to each other by a multi protein complex called cohesin. The microtubule organizing centers (centrosomes or spindle poles) help to polymerize tubulin to nucleate microtubules and begin to form the mitotic spindle and migrate to opposite ends of the

9 nucleus. At the onset of the next phase, prometaphase, the nuclear envelope undergoes breakdown during open mitosis. The microtubules at this point invade the nuclear space and attach to a specialized protein structure on the centromere called the kinetochores. During metaphase, all the chromosomes align themselves equidistant from the poles at the metaphase plate and establish bipolar attachments to the poles via kinetochore microtubules of the spindle. The bipolar attachment of every chromosome provides equal tension to maintain them at the metaphase plate. Because proper chromosome separation requires that every kinetochore be attached to microtubules in a bipolar manner, any unattached kinetochore generates a signal preventing further progression of mitosis (Chen, Waters et al. 1996). A dramatic loss of sister chromatid cohesion signals the beginning of anaphase. In early anaphase, the chromatids are initially separated and moved towards the spindle poles by shortening of the spindle microtubules where forces are exerted mainly at the kinetochores. During late anaphase, the chromatids are fully separated by the elongation, dissasembly and sliding of the spindle microtubules. At telophase, the nuclear envelope reassembles around the separated chromatids and is followed by the final step, cytokinesis which physically divides the two separated masses of DNA resulting in two identical daughter cells.

1.3.3.1. The three flavors of mitosis Higher eukaryotes undergo an open mitosis in which the nuclear envelope is completely broken down at the G2/M transition and is not reassembled until chromosomal segregation is done with in late telophase. In contrast, in lower eukaryotes like Saccharomyces cerevisiae, mitosis occurs within an intact nucleus with no disassembly of the nuclear envelope. Aspergillus nidulans however undergo a third kind of mitosis termed a partial open mitosis. In this mitosis, the nuclear pore complex (NPC) is partially disassembled to allow for rapid increase in the permeability of the nuclear envelope allowing important regulators of mitosis to enter the nucleus. It is thought that organisms that rapidly form a spindle inside the nuclear envelope utilize partial open mitosis to achieve a rapid influx of tubulin. Other organisms like yeast that maintain a

10 nuclear spindle for most of the cell cycle do not disassemble their nuclear pore complex and undergo closed mitosis (Desouza and Osmani 2007).

1.3.4. Regulation of the cell cycle 1.3.4.1. Key cell cycle discoveries Eukaryotic cells have evolved a complex network of regulatory proteins to govern progression through the cell cycle. The essence of this system is an ordered series of biochemical events that control the main steps through the cell cycle, including DNA replication and mitosis. Much of our current understanding of the cell cycle came from information gained from three main lines of investigation involving an intersection of powerful genetic screens in fungi with biochemical studies in frog and starfish oocytes. These landmark discoveries include: the identification of cyclin in starfish oocytes (Evans, Rosenthal et al. 1983), identification of cell-division-cycle mutant genes from parallel genetic screens conducted in budding yeast (Hartwell, Culotti et al. 1974), fission yeast (Nurse, Thuriaux et al. 1976) and Aspergillus nidulans (Morris 1975), and the identification and purification of maturation promoting factor (MPF)- CDK1/cyclin B from frog oocytes (Masui and Markert 1971; Gautier, Norbury et al. 1988; Lohka, Hayes et al. 1988). Subsequent sections are going to concentrate on mechanisms that regulate mitosis.

1.3.4.2. Checkpoint controls exist to monitor proper cell cycle progression An important feature of the eukaryotic cell cycle is the strict requirement of activation and completion of one phase before the cell can proceed to the next. Initial study by Leland Hartwell and Ted Weinert (Hartwell and Weinert 1989) showed that certain mutations could relieve this dependency which meant that a control mechanism existed to enforce this dependency through negative intracellular signals. These surveillance mechanisms, or so called checkpoint pathways, ensure the proper order and correct execution of cell-cycle events (Elledge 1996). Although most checkpoints are not essential for normal cell cycle progression under ideal conditions, cells with checkpoint

11 defects over time accumulate mutations due to occasional malfunctions in DNA replication, DNA repair or spindle assembly. The best understood checkpoints are the DNA structure checkpoints that arrest cells at G2/M transition in response to unreplicated or damaged DNA and the spindle assembly checkpoint (SAC) that prevents anaphase onset until all chromosomes establish a bipolar spindle attachment. Due to the fact that the ability to arrest cells in metaphase through activation of SAC is utilized in sets of experiments in this thesis, a brief overview of this checkpoint pathway follows. Sister chromatids formed during DNA replication are linked to each other by a protein complex called cohesin (Michaelis, Ciosk et al. 1997; Uhlmann and Nasmyth 1998; Ciosk, Shirayama et al. 2000; Tanaka, Fuchs et al. 2000; Haering, Lowe et al. 2002). At anaphase, there is a cascade of signaling events that allows cohesin to be degraded (Haering, Lowe et al. 2002) by the proteolytic enzyme separase (Ciosk, Zachariae et al. 1998; Marcais, Bernard et al. 2000; Zou, Stemman et al. 2002). Separase is maintained in an inactive state by its binding to securin, until all chromosomes are bipolar attached to the spindle. Once this condition is met, securin is rapidly proteolytically degraded by the APC (anaphase promoting complex) (Fang, Yu et al. 1998; Yu 2002). Loss of securin hence releases separase, freeing it to cleave cohesin and permit anaphase progression. The SAC checkpoint depends on a sensor mechanism that monitors the state of the kinetochore and tension across the kinetochore, a specialized platform on the chromosomal where microtubules attach. Unattached kinetochores are thought to function as sites of continuous binding and release of two proteins, Mad2 and Cdc20, which results in sequestering Cdc20 away from the APC preventing APC activation and arrest at metaphase(Nigg 2001). On attachment of the last kinetochore, the production of inhibitory complex Mad2-Cdc20 ceases and Cdc20 is freed to bind APC. This activates the APC and allows it to degrade securin leading to activation of anaphase.

12 1.3.4.3. Cyclin dependent kinases (CDKs) At the core of the cell cycle regulatory system is a family of protein kinases known as cyclin-dependent kinases (CDKs). The founding member of the CDK family, p34 was first identified and cloned from S. cerevisiae as cdc28 (Hartwell, Culotti et al. 1974) and later from S. pombe as cdc2 (Nurse, Thuriaux et al. 1976). Universally conserved in eukaryotes from fungi to humans (Nurse 1990), the functional conservation of this kinase was first realized when cdc28 and cdc2 could cross complement (Beach, Durkacz et al. 1982). More surprisingly, the human cdc2 homolog was cloned by complementation of the cdc2 mutation in fission yeast using a human cDNA library (Lee and Nurse 1987). The activity of cyclin-dependent kinases, including p34, rises and falls during cell cycle progression. The oscillations directly influence the phosphorylation status of intracellular proteins and lead to cyclical changes that initiate and regulate major events of the cell cycle – DNA replication, mitosis and cytokinesis. The cyclical changes of CDK activities are in turn controlled by their binding to specific regulatory subunits called cyclins. As their name implies, the kinase activities of CDKs are cyclin dependent and need binding to cyclins to have any functional kinase activity. Cyclins, first identified in marine invertebrates (Evans, Rosenthal et al. 1983) undergo cycles of synthesis and degradation. CDK levels by contrast remain constant. Oscillating changes in cyclin levels therefore result in a cycle of assembly and activation of cyclin-CDK complexes which in turn trigger specific cell cycle events. While the primary determinant of CDK activity and specificity is its binding to cyclins (cyclins directs CDK activity to specific target proteins), its activity can also be regulated by inhibitory phosphorylation of two residues in its active site (Draetta 1990; Norbury and Nurse 1992).

1.3.4.3.1. Mitotic regulation by CDK1 In S. pombe and in A.nidulans, CDK1 is required for both G1/S transition and G2/M transition (Nurse and Bissett 1981; Piggott, Rai et al. 1982; Osmani, van Peij et al. 1994). CDK1 is considered to be the key regulator of mitosis. The activation of M-CDK begins with the accumulation of the M cyclin - cyclin B during G2 and M. Although CDK1-cyclin B complexes are phosphorylated at an activating site (T161) in late S phase

13 (Morgan 1995), they are kept in an inactive state till mitosis by the action of a protein kinase, wee1 by inhibitory phosphorylations at two neighboring sites (Y15 and T14) in its active catalytic site (Gould and Nurse 1989; Krek and Nigg 1991; Den Haese, Walworth et al. 1995). Thus, by the time the cell reaches the end of G2, it contains a bulk amount of M-cyclin complexes that are ready to act, but are kept in a repressed form by the presence of two phosphate groups. What then triggers the activation of M-cyclin at the start of mitosis? CDK1 is kept in an inactive state due to the inhibitory phosphorylations by wee1 and in mitosis, is activated by the phosphatase cdc25 which removes the inhibitory phosphates and relieves the block on CDK1-cyclin B kinase activity (Draetta 1990). The characterization of CDK1 illustrates the following important points. Firstly, the cell cycle regulatory machinery is highly conserved. Next, simple model organisms can be utilized to dissect various regulatory pathways in an effective way. Finally, the use of genetics in helping us develop our understanding of the cell cycle has been crucial. The degree of conservation of the basic cell cycle machinery supports a continued notion that the study of other genes involved in regulatory pathways in such model organisms can be extended to understand higher eukaryotes. While the central role of CDK1 in promoting mitosis is beyond question, it is also clear that CDK1 activity alone is not sufficient for mitotic progression. Part of the supporting cast of regulatory proteins is made up of additional protein kinases with mitotic specific roles including Polo-like kinases (Nigg 1998), Aurora kinases (Ke, Dou et al. 2003), and the burgeoning family of NIMA-related kinases (O'Connell, Krien et al. 2003).

1.4. NIMA protein kinase Analysis of cell cycle mutants in Aspergillus nidulans has indicated that while CDK1-cyclin B kinase is essential, it is not sufficient to trigger mitosis (Osmani, McGuire et al. 1991). In the absence of the protein kinase NIMA, cells arrest in late G2 (Figure 1.6) (Osmani, Pu et al. 1988). NIMA, like CDK1, is a serine/threonine protein kinase and is required for cells to initiate mitosis. NIMA is expressed in a cell cycle dependent manner and is different from CDK1 in not requiring a binding partner for its activation. Several other genes have been identified that are also required for mitotic

14 initiation but are thought to function either directly or indirectly by affecting the kinase activity of CDK1. Remarkably, in cells arrested in G2 by conditional inactivation of NIMA, CDK1 kinase is in fact fully active (Osmani, McGuire et al. 1991). This surprising result is the only situation where CDK1 activity does not lead to mitotic entry, suggesting that NIMA plays a role in mitotic initiation that is independent of activation of CDK1 kinase activity. Furthermore, inducible expression of NIMA kinase in A.nidulans and the overexpression in S. pombe, Xenopus oocytes and human cells promotes premature meiotic and mitotic events (O'Connell, Norbury et al. 1994; Lu and Hunter 1995). In addition, expression of dominant negative forms of NIMA induces a G2 arrest in HeLa cells (Lu and Hunter 1995). Thus, NIMA kinase plays an essential role in mitotic initiation. These results provide strong evidence for the existence of NIMA-like kinases in vertebrates. Consistent with this prediction, while no true NIMA functional homologue has been identified yet outside of filamentous fungi, a number of NIMA-related kinases have been reported in both mouse and humans along with yeast (O'Connell, Krien et al. 2003).

1.4.1 Functional domains and important residues in NIMA nimA was cloned by the complementation of the nimA5 mutant phenotype and sequence analysis indicates it encodes a 79kDa protein kinase of 699 amino acids, with a catalytic domain at its N-terminal (amino acids 1-300). The catalytic domain is necessary for the kinase function of nimA as various truncated versions of the domain abrogate its catalytic activity. Two residues in the domain were shown to be important for its catalytic function, one being lysine at position 40 important for its binding to ATP and two, threonine at position 199 which needs to be phosphorylated for its activation (Figure 1.7) (Pu, Xu et al. 1995). A point mutation of either of these residues terminates NIMA kinase activity (Lu, Osmani et al. 1993; Lu and Hunter 1995). Deletion of the non-catalytic C-terminus regulatory domain does not inactivate NIMA kinase activity but does prevent the functional complementation of the temperature sensitive nimA5 mutation, showing it to be essential for function. Partial deletion of the C-terminus regulatory domain generates a highly toxic kinase even though

15 the kinase domain on its own is not toxic (Pu and Osmani 1995). This result highlights the importance of the non-catalytic regulatory domain of NIMA. While the amino- terminus encodes the functional kinase domain, the carboxy-terminus is critical for regulating this catalytic function. Several functional motifs within this region constitute these overlapping regulatory systems. First, the timely proteolytic degradation of NIMA is essential for normal cell cycle progression. Although, the C-terminal region of NIMA is not essential for kinase function, it is believed to be important in the proteolytic degradation of the protein (O'Connell, Norbury et al. 1994; Pu and Osmani 1995). Sequences rich in proline (P), glutamic acid (E), serine (S), and threonine (T) are present in this region, so called PEST sequences. PEST sequences are believed to be involved in facilitating rapid degradation of the proteins in which they are contained (Rechsteiner and Rogers 1996; Roth and Davis 2000). Lack of these PEST sequences produces a version of NIMA which is resistant to degradation and therefore far more stable than normal NIMA (Pu and Osmani 1995; O'Connell, Krien et al. 2003). Expression of these stable forms of NIMA is highly toxic and is capable of inducing chromosome condensation in higher eukaryotes. Additionally, expression of these forms of NIMA blocks mitotic exit (Pu and Osmani 1995). Three nuclear localization sequences are also present. Two of the nuclear localization signals are found in the non-catalytic domain, and the other is located at the border of the two functional domains. Finally, two coiled-coil domains exist just outside of the catalytic domain. Coiled-coil motifs are present in a broad array of proteins and are believed to be involved in protein-protein interactions. Coiled coils are alpha-helical structural motifs comprised of a seven amino acid repeat pattern. Residues in the first and fourth positions are hydrophobic while residues in the fifth and seventh positions are polar or charged. These motifs are thought to play a role in protein-protein interactions by associating with similar motifs in other proteins. Some studies have provided evidence that coiled-coil domains may participate directly in homodimer and heterodimer formation (Blake, Tinsley et al. 1995; Chiu, Revenkova et al. 2004; Nikolay, Wiederkehr et al. 2004) and therefore, the coiled-coil domains of NIMA may allow for protein-protein interaction of NIMA with other cell cycle regulatory proteins. It has been demonstrated that

16 overexpression of the non-catalytic regulatory terminus of NIMA produces the same phenotype, that of an arrest in late G2, as overexpression of kinase inactive NIMA (Lu and Means 1994). This dominant negative effect also suggests that the carboxy-terminus of NIMA, potentially the coiled coil regions, is the site of protein-protein interactions and that the mutant version out competes essential binding substrates away from wild type NIMA causing cell cycle progression defects.

1.4.2. NIMA levels through the cell cycle Previous studies have shown that the mRNA levels and protein levels of nimA and the protein kinase activity of NIMA is cell cycle regulated (Figure 1.8) (Osmani, May et al. 1987; Osmani, McGuire et al. 1991; Osmani, O'Donnell et al. 1991). NIMA levels remain low through out S phase. It begins to accumulate starting in G2 and reaches its maximal level at M. The level of NIMA protein kinase activity assayed from mitotic cells is 20 fold higher than that measured in cells arrested in S phase (Ye, Xu et al. 1995). As nuclear division is completed, NIMA protein and kinase activity are reduced. It is not completely understood if the variation in nimA transcript levels is due to changes in the rate of transcription from the gene or due to changes in the rate of turnover of the mRNA. However, the fact that nimA transcript levels fall rapidly during exit from mitosis suggests that the stability of nimA transcripts is important in controlling nimA mRNA levels. During interphase phosphorylated, but not fully activated, forms of NIMA accumulate. These forms continue to accumulate in the absence of active CDK1 (Ye, Xu et al. 1995). Upon entry into mitosis, fully activated NIMA is targeted for degradation (Pu and Osmani 1995). It has been shown that NIMA is highly destabilized during an extended S phase following addition of hydroxyurea and that this destabilization is dependant on the APC component BIME (Osmani and Ye 1996). It follows that the APC should also target NIMA kinase for destruction to allow for mitotic exit. This idea is supported by the fact that an APC mutant (bimE7) can override a nimA5 temperature sensitive G2 arrest (cells here have elevated NIMA levels but are not completely activated) and partially circumvent the requirement for NIMA activation in the process

17 of mitotic initiation (Osmani, O'Donnell et al. 1991). It has been proposed that the hyperphosphorylation of NIMA at the G2 to M transition may act as a signal for proteolysis, effectively recruiting the APC, to trigger NIMA degradation (Ye, Xu et al. 1995). Additionally, the PEST sequences almost certainly play a critical role in the degradation of NIMA during mitosis as versions of NIMA lacking these sequences are highly stabilized (Pu and Osmani, 1995a; O'Connell et al., 1994).

1.4.3. NIMA protein kinase is hyperphosphorylated during mitosis Cellular NIMA exists as a phosphoprotein and bacterially expressed NIMA is also phosphorylated on multiple serine/threonine residues. Further studies reveal that these phosphorylations in the active site called T-loop are essential for the NIMA kinase activity as the T loop region resides just prior to the proposed activating phosphorylation site of NIMA (Lu, Osmani et al. 1993). When NIMA is dephosphorylated, it becomes inactive. During interphase NIMA is initially unphosphorylated and hence is inactive as a kinase. Later in G2 NIMA is phosphorylated and shows basal level of kinase activity. Autophosphorylation of NIMA may be one means by which NIMA is phosphorylated and activated. Another possibility is that NIMA may well be an intermediate protein in a cascade of protein kinases. Inactivation of cdc25 function prevents activation of CDK1- cyclin B by tyrosine phosphorylation resulting in a G2 arrest. In the absence of cdc25 function, NIMA protein accumulates in a phosphorylated state but is only partially active. Release from the G2 arrest causes activation of CDK1-cyclin B and this result in the further increase in the kinase activity of NIMA and causes full hyperphosphorylation of NIMA (Ye, Xu et al. 1995). It is this hyperphosphorylated form of NIMA which is present during mitosis, and which is rapidly degraded during exit from mitosis. This fact confines maximal NIMA activity to a defined period of early mitosis.

18 1.4. 4. NIMA interacting proteins and targets

In the recent past, the identification of various NIMA interacting proteins has helped to further define the role of NIMA at mitotic entry. The identification of two extragenic suppressors of the nimA1 mutant – sonA1 and sonB1 show them to be nuclear pore complex proteins that physically interact with each other. The specific isolation of these two nucleoporins as NIMA interacting proteins highlights the important role played by the nuclear pore complex at the G2/M transition in localizing important mitotic regulators to the nucleus. The sonA1 mutation, a homolog of yeast Gle2/Rae2, suppresses the predominantly cytoplasmic accumulation of CDK1/cyclin B phenotype of nimA1 arrested cells and allows for nuclear accumulation of CDK1/cyclinB promoting entry into mitosis (De Souza, Horn et al. 2003; De Souza, Osmani et al. 2004). On the other hand, the sonB1 mutation is a point mutation in its sonA interacting domain resulting in the weakening of the SONA - SONB interaction. This further leads to accumulation of NIMA1 protein into the nucleus in nimA1 arrested cells. Recent data suggests that in wild type cells, NIMA phosphorylates SONB (this may lead to weakening of its interaction with SONA, similar to the sonB1 mutation) and disperses it from the nuclear pore complex at mitotic entry. Hence, NIMA controls mitotic onset by regulating the localization of mitosis specific regulators, including CDK1/cyclinB, through interaction with the nuclear pore complex (Wu, Osmani et al. 1998; De Souza, Horn et al. 2003; De Souza, Osmani et al. 2004). In a separate screen, using the yeast two-hybrid system, two other nimA interacting genes were identified. Termed for two-hybrid interactors of NIMA, tinA and tinC, helped to elucidate the role played by NIMA at the spindle pole and in nuclear membrane dynamics, respectively (Osmani, Davies et al. 2003; Davies, Osmani et al. 2004). The tinA gene encodes for a protein that is cell cycle regulated and physically interacts with NIMA, specifically at G2. The finding that TINA localizes to the spindle pole bodies during mitosis in a microtubule dependent manner and negatively regulates the nucleation of astral microtubules while requiring active NIMA suggests that TINA may help in aiding NIMA functions at the spindle poles in forming the mitotic intranuclear spindle (Osmani, Davies et al. 2003). Because NIMA localizes to the spindle 19 and the spindle pole bodies during mitosis, and is required for spindle formation, it likely plays a role in spindle formation. The nimA related kinase in S. pombe, Fin1p has also been shown to a function in spindle formation (Grallert and Hagan 2002) and therefore the role of NIMA in regulating spindles is a conserved one. The tinC gene product localizes to the cytoplasm instead and also physically interacts with NIMA in a cell cycle dependent manner (Osmani, Davies et al. 2003). Expression of amino terminal truncated version of TINC causes multiple rounds of defective mitosis and a premature disappearance of NIMA protein from the cells. There is an accumulation of the truncated version of TINC into membranous bodies and an uncoupling of DNA segregation from nuclear membrane fission during mitotic exit (Davies, Osmani et al. 2004). This suggests that the process of separation of the nuclear envelope into two daughter nuclei is a regulated process in A.nidulans. Given the interaction between TINC and NIMA and the apparent destabilization of NIMA in the presence of the truncated version of TINC leading to defective membrane dynamics, there is a possibility that TINC helps in stabilizing NIMA during mitosis and suggests a role for NIMA in nuclear envelope fission at mitotic exit.

1.4.5. NIMA related kinases and their role in the cell cycle The identification of nimA and the characterization of its essential mitotic specific function in initiated a search for NIMA homologues in other organisms. To date the only true functional homolog of NIMA capable of complementing a nimA (nimA5) mutation is nim-1 from N. crassa (Pu, Xu et al. 1995). Nim-1 was isolated by a low stringency hybridization screening of a N. crassa genomic library. Hybridizing fragments were isolated and were tested for their ability to complement nimA5 (Pu, Xu et al. 1995). In the beginning, it appeared that NIMA function might only be required for nuclear division events in the syncitial filamentous type fungi as no true homologs of NIMA were found in other organisms. Later, a single NIMA-related kinase was identified in fission and budding yeast (Fin1p and KIN3 respectively). Fin1 was also identified in a genetic screen for mutations affecting spindle formation (Grallert and Hagan 2002). As noted previously, the carboxy terminus of NIMA is characterized by the presence of 20 coiled-coil and PEST domains. Many of the fungal NIMA-related kinases contain similar domains in their carboxy-termini, while some homologues contain additional protein domains (O'Connell, Krien et al. 2003). New data also emerged in the mid 1990s, showing that ectopic expression of Aspergillus NIMA in metazoan cells could induce hallmarks of mitosis, most notable DNA condensation and nuclear envelope breakdown (O'Connell, Norbury et al. 1994; Lu and Hunter 1995). These results were the first evidence that, like other key regulators of the cell cycle, kinases related to NIMA may be important mitotic regulators in higher eukaryotes as well. The first mammalian NIMA-related kinases (Neks), Nek1, Nek2 and Nek3 were first identified in the early 1990s (Letwin, Mizzen et al. 1992; Schultz and Nigg 1993). However, with the completion of the human genome and murine genome sequence unexpectedly revealed the presence of 11 genes that encoded a distinct family of mammalian serine/threonine kinases related to NIMA (Figure 1.9) (Forrest, Taylor et al. 2003). Termed Nek1 to Nek11, they constitute approximately 2% of the entire human kinome (O'Regan, Blot et al. 2007) and share approximately 40-45% identity with NIMA within their N-terminal catalytic domain, though the C-terminal non catalytic domain are highly divergent suggesting that each kinase might have a distinct function (O'Connell, Krien et al. 2003). Nevertheless, new data now suggests that at least 4 of these Neks (Nek2, Nek6, Nek7 and Nek9) are likely to be important regulators of mitotic progression. They have been shown to localize to the microtubule organizing centers (equivalent of the spindle pole bodies) where they have important function in microtubule organization and mitotic spindle formation. The importance of NIMA function the G2-M transition suggested that NIMA-related proteins would also act at this point in the cell cycle. Detailed investigation of some of the NIMA-related kinases has identified mitotic specific roles. The S. pombe NIMA homologue termed Fin1 is capable of inducing chromatin condensation in the absence of CDK1 activity and undergoes cell cycle mediated fluctuations (Krien, Bugg et al. 1998), however, Fin1p levels appear to peak at the metaphase to anaphase transition, a point later in mitosis than NIMA (Krien, West et al.

21 2002). Additionally, loss of fin1 does produce severe nuclear envelope perturbations and causes synthetic lethality with mutants of the mitotic spindle checkpoint (Krien, Bugg et al. 1998; Krien, West et al. 2002). Additional ts fin1 mutants displayed defects in spindle pole body microtubule nucleation and failure to recruit Polo kinase to the spindle pole body (Grallert and Hagan 2002). Finally, a potential role for Fin1p in regulating spindle pole body maturation has been suggested. It is proposed that Fin1p modulates activity of the SIN (Septation Initiation Network) pathway by having specific functions in the process of maturation of the spindle pole (Grallert, Krapp et al. 2004). Of the eleven mammalian Neks, the most closely related by sequence within the catalytic domain to NIMA and Fin1 is Nek2 and, biochemically, Nek2 and NIMA share many common properties (Lu, Osmani et al. 1993; Fry, Schultz et al. 1995). Like NIMA and Fin1, Nek2 exhibits a cell cycle dependent expression and protein profile though Nek2 does not rescue the nimA5 mutation or Fin1 deletion. Nek2 appears to play a direct role in enabling bipolar spindle formation through initiating the separation of centrosomes at the G2/M transition (Fry, Meraldi et al. 1998). C-Nap1 has been identified as a Nek2 target with a role in centrosome cohesion and establishing a mitotic spindle (Fry, Mayor et al. 1998). An additional Nek2 target, Hec1, binds to kinetochores and is involved in the spindle checkpoint and regulating kinetochore attachments through its interactions with SMC proteins and centromere protein Ctf19p (Chen, Riley et al. 2002; Martin-Lluesma, Stucke et al. 2002; DeLuca, Howell et al. 2003; Hori, Haraguchi et al. 2003). Kin3, the S. cerevisiae NIMA related kinase has also been shown to interact with Hec1 (Chen, Riley et al. 2002). While cells lacking Kin3 are able to survive, the expression of a version of Kin3 equivalent to nimA7 mutation results in severe defects in chromosome segregation (Chen, Riley et al. 2002). More recently a study has identified a direct interaction between the spindle checkpoint component MAD1 and Nek2 (Lou, Yao et al. 2004). In addition to Nek2 two additional NIMA related kinases, Nek6 and Nek9, have potential mitotic specific regulatory roles (Kandli, Feige et al. 2000; Hashimoto, Akita et al. 2002; Holland, Milne et al. 2002; Roig, Mikhailov et al. 2002; Yin, Shao et al. 2003). Nek 6 is phosphorylated and activated at mitosis (Yin, Shao et al. 2003). Expression of

22 kinase negative forms of Nek6 results in a metaphase arrest followed by apoptosis (Yin, Shao et al. 2003). Nek 9 is also phosphorylated and activated at mitosis (Roig, Mikhailov et al. 2002). Nek9 contains an RCC1 homology domain and interacts with the Ran GTPase, a regulator of diverse functions including nucleocytoplasmic transport, spindle dynamics and nuclear envelope reassembly (Roig, Mikhailov et al. 2002). Interference with Nek9 function by antibody injection produces chromosome segregation defects. Finally, Nek 9 binds to and activates Nek6 (Roig, Mikhailov et al. 2002; Yin, Shao et al. 2003). Certain members of the mammalian family of Neks have also been implicated in functions apart from mitosis. The NIMA related kinase Nek11 has been shown to act as a DNA replication/damage stress responsive kinase (Noguchi, Fukazawa et al. 2002). Additionally, a novel interaction between Nek11 and Nek2 has been discovered. This interaction occurs in the nucleolus and is specific to the phosphorylation status of Nek2. While phosphorylated Nek2 binds Nek11, a kinase inactive Nek2 fails to do so (Noguchi, Fukazawa et al. 2004). Moreover, Nek2 directly phosphorylates Nek11 and is responsible for its activation during the G1/S phase (Noguchi, Fukazawa et al. 2004). Therefore, Nek11 activation by Nek2 may represent a novel function of the NIMA related kinases in the nucleolus.

1.5. The Nucleolus The nucleolus has attracted the attention of investigators for decades due to its ubiquitous presence in the eukaryotic nucleus and its core role in cellular physiology. The nucleolus is a membraneless intranuclear organelle and exists in various numbers, sizes and shapes within the nuclei of different plant and animal cells. When viewed under the electron microscope, it appears as non-uniformly stained bodies in higher eukaryotes and as a single crescent shaped structure in the yeast, Saccharomyces cerevisiae. The nucleolus occupies almost one-third to on-half of the nucleus in yeast (Warner 1990). In the early 1930s, Heitz and McClintock, followed by many other cytologists, firmly established nucleoli to be directly involved with chromosomal activity (Heitz 1931; Heitz 1933; McClintock 1934). Later, two principle discoveries in the 1960s allowed for the 23 identification of the nucleolus as the cellular center for ribosome biosynthesis in eukaryotes (Perry and Errera 1961; Perry 1962; Brown and Gurdon 1964; Birnstiel, Wallace et al. 1966; Ritossa, Atwood et al. 1966) . Thus the major function associated with the nucleolus is ribosome biogenesis involving transcription of ribosomal genes, processing of ribosomal transcripts and subsequent assembly of ribosomes. Because the production of ribosomes is a major metabolic activity, the function of the nucleolus is tightly linked to cell growth and proliferation. Over the past few years, the ability to purify the nucleolus on a large scale combined with important advances in the identification and analysis of proteins using mass spectrometry, has revealed new functions of the nucleolus including gene silencing, cell cycle progression, senescence, cellular aging, stress response, mRNA processing and quality control of various ribonucleoproteins involved in protein synthesis (Johnson, Marciniak et al. 1998; Pederson 1998; Cockell and Gasser 1999; Straight, Shou et al. 1999; Visintin, Hwang et al. 1999; Zhang and Xiong 1999; Pederson and Politz 2000; Visintin and Amon 2000). The purpose of this section is to provide an integrated view of our present day knowledge of the nucleolus as the center of ribosomal biogenesis and discuss recent progress in our understanding of nucleolar architecture and its relationship to nucleolar function. A brief introduction to the cell cycle regulation of the nucleolus will be given here but a more detailed discussion will follow in chapter 7.

1.5.1. Repeated copies of the ribosomal RNA genes are the organizers of the nucleolus The ribosome is a complex molecular structure comprised of two distinct components, the large and the small subunit, that fit together and work as one to help translate mRNAs into polypeptide chains during protein synthesis. Eukaryotes have 80S ribosomes, each consisting of a 60S (large) and a 40S (small) subunit. The two components are made and present in the cell in equimolar amounts and are structures built from four distinct rRNAs and about 80 proteins. The genes involved in the biogenesis of ribosomes include the rRNA genes for the four rRNAs (L-rRNA, S-rRNA, 5.8S rRNA and 5S rRNA) and the genes required for the expression of structural proteins

24 of the ribosome, r-proteins. Additionally, they include the genes for the specific enzymes and proteins specialized in transcription, modification and processing mechanisms resulting in the formation of mature ribosomes. In all eukaryotes, the genes for L-rRNA, S-rRNA and 5.8S rRNA exist adjacent to each other and are initially transcribed as a large precursor RNA molecule, which may be thought of a single transcription unit. The precursor is then converted in the nucleoli, via intermediates, to the mature forms of ribosomal RNAs. All four rRNA genes are highly repeated; each being represented 102 to 103 copies per haploid genome. The multiple sets of rRNA transcription units are arranged in tandem separated by non transcribed spacer sequences into one or several clusters along chromosomes. During mitosis, these clusters of genes are seen as secondary constrictions in the respective chromosomes. The rDNA repeats are termed as “nucleolar organizer regions” (NOR) because they possess the capacity to form nucleoli at telophase. Secondary constrictions are distinct from primary constrictions (often called centromeres or kinetochores), the attachment site to the mitotic spindle. NOR were first identified in the corn Zea mays when deletion of the secondary constriction resulted in homozygous anucleolate mutants incapable of nucleolus formation (McClintock 1934). Further studies using mutants with variable number of NOR from the fruit fly Drosophila melanogaster (Ritossa and Spiegelman 1965) and the frog Xenopus laevis (Birnstiel, Wallace et al. 1966) proved NOR to be the chromosomal site of clustered rRNA genes. Mutants with deleted NOR resulted in complete abolishment of rRNA synthesis. Direct proof for the location of the rRNA genes in the NOR was obtained by in situ rRNA: DNA hybridization (Gall and Pardue 1969; John, Birnstiel et al. 1969). The numbers of nucleolar organizers vary from one organism to another and the number of rRNA genes in a given NOR may vary within a particular species. Such variation may be an inevitable consequence of the tandem arrangement of multiple rRNA genes. It has been proposed that tandemly arrayed identical sequences should result in unequal crossing-over events during mitosis (sister chromatid exchanges) or meiosis (sister and non-sister chromatid exchanges) (Tartof 1973; Smith 1974; Tartof 1975). Moreover, the occurrence of unequal crossing over in the S. cerevisiae array of rRNA

25 genes in sister chromatids was shown to take place during mitosis and meiosis(Petes 1980; Szostak and Wu 1980) These data support the idea that unequal crossing over at the clusters of rRNA genes may be a major factor in determining multiplicity of the rRNA genes.

1.5.2. Chromosomal location of repeated copies of ribosomal genes Clustering of rRNA genes on the chromosome is a typical feature of prokaryotes (Nomura and Morgan 1977) and this pattern is preserved in eukaryotes. Lower eukaryotes like Saccharomyces cerevisiae contain 100-120 rRNA genes per haploid genome (Retel and Planta 1968; Schweizer, MacKechnie et al. 1969) where 90% of the rRNA genes are confined to a single chromosome (Petes and Botstein 1977). The chromosomal location of rRNA gene clusters in a broad variety of higher eukaryotes has also been established. Generally, one to six chromosomes carry the NORs (Smetana, Gyorkey et al. 1970; Lima-de-Faria 1980). Eukaryotes are also capable of differential replication of their rRNA genes through formation of extrachromosomal rDNA repeating units that may be maintained extrachromosomally or might subsequently reintegrate into the genome. As a result in some cases the cell can have more or fewer rRNA genes which do not correspond strictly to the ploidy of the cell. Generally, differential replication takes place in order to meet unusually high demands for ribosome production made on the cell. In some cells and organisms differential replication of rRNA genes tales places as a correction mechanism, aimed at keeping the number of rRNA genes above a critical level, when they are reduced by unequal crossing over or mutational events (Macgregor 1972; Spear 1974; Macgregor and del Pino 1982).

1.5.3. Ribosomal gene transcription is required for formation of the nucleolus The initial formation of the nucleolus requires the active transcription of the rRNA genes by RNA polymerase I (Karpen, Schaefer et al. 1988; Scheer and Weisenberger 1994). In situ hybridization studies of interphase nuclei have shown the nucleolus to contain rDNA (Gall and Pardue 1969; John, Birnstiel et al. 1969) while the insertion of an rRNA gene into the euchromatic region of a chromosome also results in

26 formation of a nucleolus at that site (Karpen, Schaefer et al. 1988). Transcription of ribosomal genes for ribosome biogenesis thus is responsible for the formation of the nucleolus. Note though that a significant fraction, often >50%, of rRNA genes remain transcriptionally silent and these NORs are thought not to form a nucleolar structure (Pardue and Gall 1970; Batistoni, Andronico et al. 1978) suggesting a regulatory mechanism for turning on only certain clusters of rRNA genes. Transcription is catalyzed by various different DNA dependent RNA polymerases. In eukaryotes, three types of RNA polymerases exist which are specialized in the transcription of different sets of genes: RNA polymerase I for rRNA genes (Grummt 1999), II for r-protein (Kornberg 1999), and other genes; and III for 5S rRNA genes (Willis 1993). Multiple RNA polymerase I complexes transcribe in tandem each rRNA transcription unit. This process can be visualized by chromosome spreading (Miller and Beatty 1969; Miller and Beatty 1969) and observation of the exposed transcription complexes. Active transcription units (pre-ribosomes) are visualized as comprising axes of rDNA covered with densely packed lateral fibrils forming a gradient of increasing length (Ghosh and Paweletz 1996). The growing rRNA is simultaneously coated with proteins to form the ribosomal nucleoprotein particles (Chooi and Leiby 1981). The estimated elongation rate in vivo of rRNA transcription units is in the range of 20-40 nucleotides per second. The assembly of pre-ribosomes starts during transcription of rRNA genes itself and the growing rRNA chain is automatically coated with ribosomal and non-ribosomal proteins. Thus, assembly of pre-ribosomes takes place simultaneously with transcription to release upon termination an already defined particle. The maturation of the primary preribosomes is a complex, sequential process. Maturation of pre-rRNA in eukaryotes starts after transcription termination and release of the primary pre-ribosomal nucleoprotein particle from the rDNA template. Maturation involves several endonuclease cleavages of the pre-rRNA and the addition of many r- proteins resulting in the formation of nascent small and large ribosomal particle. Almost all maturation steps takes place in the nucleolus and end with the release of nascent particles and their transport into the cytoplasm.

27 1.5.4. Molecular architecture of the Nucleolus Three main specific components can be identified in most eukaryotic nucleoli when stained with uranyl-acetate and visualized by transmission electron microscopy. Named according to their appearance, the central area is a lightly stained region called the fibrillar center (FC). Surrounding the fibrillar center is a densely stained and highly contrasted region called the dense fibrillar component (DFC). The fibrillar regions are embedded and surrounded by the granular component (GC), mainly composed of granules of 15-20nm in diameter. The fibrillar center form discrete structures and are connected by a network of dense fibrillar components (Junera, Masson et al. 1995). Nucleoli of different cell types exhibit a variable number of fibrillar centers of variable sizes, the size being inversely proportionate to the number (Pebusque and Seite 1981; Hozak, Novak et al. 1989). Many eukaryotes have a bipartite nucleolus made of only fibrillarin center and granular component. A recent proposition suggests that during evolution, a third nucleolar compartment emerged at the transition between the anamniotes and the amniotes, following a substantial increase in size of the rDNA intergenic region (Thiry and Lafontaine 2005). This conclusion has an important implication for understanding the structure-function relationships between rDNA and the nucleolus. General mapping of ribosome biogenesis in these morphologically distinct regions of the nucleolus has been followed which suggest the following model. Transcription of rRNA genes occurs either in the periphery of the fibrillarin center or at the border of the fibrillarin center and dense fibrillarin component. The nascent transcripts accumulate in the dense fibrillarin center (Hozak, Cook et al. 1994; Cmarko, Verschure et al. 2000). The pre-ribosomal RNA transcripts are modified in the dense fibrillarin center by small nucleolar ribonucleoproteins (Cmarko, Verschure et al. 2000). The final maturation of the pre-ribosomal ribonucleoprotein, and assembly with ribosomal proteins, occurs mostly in the granular center region. Not surprisingly, the distribution of the machineries involved in each step closely correlates with the region of their function. The fibrillarin center contains the tandemly repeated rRNA genes and proteins essential for the transcription process such as RNA polymerase I, DNA

28 topisomerase I and the transcription factor UBF (Thiry and Goessens 1992; Schwarzacher and Wachtler 1993; Hannan, Hannan et al. 1998). Proteins like fibrillarin (Hozak 1995) and nucleolin (Ma, Matsunaga et al. 2007) that are involved in early stages of rRNA processing, localize in the dense fibrillarin component while proteins that are involved in the intermediate or later stages of ribosome biogenesis, such as B23 and Nop52 (Biggiogera, Burki et al. 1990; Angelier, Tramier et al. 2005), localize to the granular component. These findings have led to the general hypothesis that specific compartments of the nucleolus have specific functions.

1.5.5. Nucleolar constituents are highly dynamic Transcriptional arrest of RNA polymerases I and II has been shown to be accompanied by the sorting and rearranging of nuclear proteins and RNAs into defined nuclear sub-domains. Transcriptional inactivation of rRNA genes leads to the segregation of the three defining regions of the nucleolus (FC, DFC, GC) into three distinct but juxtaposed domains that retain many of their original protein and RNA components. This is accompanied by the release of several GC proteins into the nucleoplasm and the influx of a significant number of nucleoplasmic proteins, many of which are RNA binding proteins, into large nucleolar caps. The nucleolus is thus sensitive to the transcriptional profile of the cell, and the status of transcriptional activity is reflected in nucleolar structure. The extent of nuclear, nucleolar and RNA dynamics has been completely reevaluated by the ability to track movements of individual proteins by fluorescence tagging and by photo bleaching experiments that measure intracellular mobility and residence time of proteins (Chen, Dundr et al. 2005; Olson and Dundr 2005; Shav-Tal, Blechman et al. 2005; McDonald, Carrero et al. 2006). Current results show that there is free diffusion of proteins through the nucleus, including the nucleolus (Handwerger, Cordero et al. 2005), and that the mean residence time for nucleolar proteins in the nucleolus is a few tenths of a second (Raska, Shaw et al. 2006). The nucleolus exists as a distinct structure because certain proteins, some of which still remain uncharacterized, have binding affinities to rDNA, resulting in the formation of a stable core which then

29 acts as a building core structure for other nucleolar protein complex interactions. The surrounding nucleoplasmic components continually exchange with this core structure (Raska, Shaw et al. 2006). The steady state composition for the nucleolus therefore is a result of the nucleolar residence time of non-nucleolar proteins, which do not find binding partners in the nucleoli and those of nucleolar proteins that spend a longer time at the nucleolus. The nucleolus thus represents a dynamic structure in constant equilibrium with the surrounding nucleoplasm (Chen, Dundr et al. 2005; Handwerger, Cordero et al. 2005; Olson and Dundr 2005; Shav-Tal, Blechman et al. 2005; Raska, Shaw et al. 2006). It is important to note that nucleolar proteins do not possess a common nucleolar localization signal which allows their recruitment to the nucleolus. It is their functional interaction with other molecules already present in the nucleolus that instead acts as a retention signal (Misteli 2005). More recently, a GTP switch is suggested to be involved in the regulation of protein targeting to the nucleolus and to be responsive to extracellular stimuli via signaling pathways (Misteli 2005; Tsai and McKay 2005).

1.5.6. Nucleolar proteomic studies: new information regarding nucleolar content The ability to isolate and purify large amounts of nucleoli that retain transcriptional activity to some extent (Cheutin, O'Donohue et al. 2002; Lam, Trinkle- Mulcahy et al. 2005), prompted the recent study and analysis of nucleolar protein composition in human and plant cells, using high-throughput mass spectrometry based techniques. These studies have identified more than 200 plant proteins and over 700 human proteins that stably co-purify with the nucleolus (Andersen, Lyon et al. 2002; Scherl, Coute et al. 2002; Andersen, Lam et al. 2005; Pendle, Clark et al. 2005). More than 90% of the human nucleolar proteins identified from the study showed clear homologues in yeast (Andersen, Lam et al. 2005) demonstrating the conservation of nucleolar proteins through evolution. Bioinformatics analyses of the proteomic data have suggested potential functions for ~150 previously uncharacterized nucleolar proteins (Heix, Vente et al. 1998; Leung, Andersen et al. 2003; Coute, Burgess et al. 2006; Vollmer, Horth et al. 2006). Many proteins related to cell cycle regulation (~3.5%), DNA damage repair (~1%) and pre

30 mRNA processing (~5%) have been detected (Lam, Trinkle-Mulcahy et al. 2005) and are suggestive of additional roles of the nucleolus beyond ribosomal biogenesis. Other cellular activities linked to the nucleolus include RNA editing (Sansam, Wells et al. 2003), DNA damage repair (Sansam, Wells et al. 2003), telomere metabolism (Kieffer- Kwon, Martianov et al. 2004; Zhang, Hemmerich et al. 2004), tRNA processing (Paushkin, Patel et al. 2004) and regulation of protein stability (Mekhail, Gunaratnam et al. 2004). A separate study undertaken by Andersen et al. allowed a quantitative analysis of the human nucleoli proteome. In vivo fluorescent imaging techniques revealing protein kinetics were directly compared to endogenous protein changes measured by proteomics. Under different metabolic conditions that affect nucleolar morphology, the study showed that stably associated nucleolar proteins, such as RNA polymerase I, exited from or accumulated in the nucleolus with similar kinetics, whereas protein components of the large and small ribosomal subunits left the nucleolus with markedly changed kinetics (Andersen, Lam et al. 2005). This approach allowed for the temporal characterization of protein flux through the nucleolus and demonstrated that the nucleolar proteome can significantly change in response to changes in growth conditions.

1.5.7. Maintenance of nucleolar structure can be uncoupled from ribosomal RNA transcription Regarding the question of what normally maintains the shape and structure of the nucleolus, it has been proposed that RNA polymerase I driven transcription, with ribosomal genes serving as the seeding site, is sufficient to organize and maintain the nucleolus. However, experiments performed with Xenopus germ cell proteins, FRGY2a and FRGY2b, has shown that the nucleolar structure can be reversibly disassembled in vivo and in vitro even in the presence of ongoing ribosomal RNA transcription (Gonda, Fowler et al. 2003). The study proposes that these proteins may sequester B23, a major nucleolar phosphoprotein, resulting in the uncoupling of nucleolar morphology from Pol I driven transcription. Therefore, although ribosomal gene transcription by Pol I is required for the initial assembly of nucleolar structure, it is not sufficient for its maintenance.

31 1.5.8. Nucleolar structure during mitosis In yeast, the nucleolus and its associated proteins remains intact during mitosis, ultimately being separated along the mitotic spindles during late stages of division (Granot and Snyder 1991; Fuchs and Loidl 2004). In contrast, the nucleolus from higher eukaryotes disassembles during mitosis and various nucleolar proteins leave the structure in an apparently orderly progression (Rabut, Lenart et al. 2004). The granular component disappears first, followed by the dense fibrillar component. At the same time, the NORs become visible on chromosomes. Proteins from these two regions localize to the perichromosomal areas and remain there till the end of mitosis, at which point small round structures called pre-nucleolar bodies (PNBs) containing nucleolar proteins begin to appear in the daughter nucleus. Initiation of transcription by RNA polymerase I then triggers the recruitment of PNBs to the NORs and establishes a complete nucleolus (Scheer and Weisenberger 1994). There are therefore fundamental differences between the mechanisms of mitotic nucleolar segregation between lower and higher eukaryotes. These differences likely relate to the fact that unicellular yeasts, such as S. cerevisiae and S. pombe, undergo closed mitoses within intact nuclei whereas higher eukaryotes undergo open mitosis during which the nucleus is disassembled (Rabut, Lenart et al. 2004). Thus, our current understanding of the functional organization of the nucleolus is that it is a self-organizing, dynamic system (Misteli 2001). Nucleolar components are continuously exchanged with the nucleoplasm while reformation of the nucleolus is coupled to transcription of the rRNA genes by RNA pol I. It is still unclear what components provide the structural integrity of the nucleolus. It is also not known how nucleolar assembly and disassembly is regulated during mitosis and what may be the full range of function for the nucleolar proteins newly identified by proteomic studies. Therefore, much needs to be learned concerning the nucleolus. However, chapter 7 in this thesis will specifically deal with the mitotic regulation of the nucleolus in Aspergillus nidulans and show it to unexpectedly undergo similar nucleolar dynamics to human nucleoli.

32 Conidiospore

A C Conidiospore

Secondary sterigmata Primary sterigmata

Vesicle B

Foot cell

Germling

Figure 1.1. Asexual development of A. nidulans (A) The conidiospore is a uninucleate haploid spore. (B) Under favorable growth conditions, the spore undergoes polarized growth and rounds of nuclear division to produce a multi-nucleate germling. (C) After a period of growth specialized cells termed foot cells develop within the hyphae. An aerial hyphal branch emerges from the foot cell and the tip swells to form a vesicle. Primary and secondary sterigmata develop from the vesicle. Mitotic division and cytokinesis at the apex of the secondary sterigmata produces chains of conidia.

33

(Reprinted by permission from Macmillan Publishers Ltd: Nature Reviews Genetics, Vol. 3, Casselton, L. and Zolan, M., The art and design of genetic screens: filamentous fungi, pp. 683-697, Copyright 2002)

Figure 1.2. Lifecycles of A. nidulans A. nidulans is a filamentous fungus which is capable of undergoing both an asexual and sexual lifecycle. A. nidulans exists primarily in a haploid state although the organism will form transient diploid nuclei during the sexual cycle in specialized cells termed asci. In a laboratory setting diploids are stable and can be readily maintained on limiting media. 34

(Image courtesy of Dr. N. Ronald Morris)

Figure 1.3. Spore color mutants of A. nidulans The image depicts a plate of A. nidulans colonies representing the range of color mutations available. Wild type isolates of A. nidulans are green in color while other colors like chartreuse, fawn, white, and yellow, are produced by mutations in genes regulating spore color. These color mutations provide useful tools during genetic manipulations.

35 Polarized Growth Mitotic Nuclear Division Mitotic Nuclear

(Image courtesy Dr. Stephen A. Osmani)

Figure 1.4. Polarized growth and nuclear division in A. nidulans A. nidulans is a multi-nucleate organism. During germination the A. nidulans spore undergoes initial isotropic expansion followed by highly polarized growth with synchronous rounds of nuclear division. The number of mitoses occurred in a cell can be determined by counting the number of nuclei. For example, the longest cell pictured last has undergone four rounds of mitosis.

36 M

G2 M PHASE

G1

I NT E S ERPHAS

(Courtesy Jon Davies)

Figure 1.5. The eukaryotic cell cycle The eukaryotic cell cycle is composed of interphase and M phase, with interphase occupying the vast majority of the cell cycle. Interphase can be further dissected into two

Gap phases (G1 and G2) and DNA replication termed S phase. M phase consists of mitosis followed by cytokinesis. Cells proceed sequentially through the cell cycle from G1 to S to G2 to M, with progression into the next phase requiring successful completion of the preceding phase.

37

Figure 1.6. nim mutants Wild type strains of A. nidulans undergo rounds of nuclear division with polarized growth to generate a multi-nucleate organism. NIMA activity is required for each round of mitosis. In the absence of functional NIMA, cells fail to undergo nuclear division and arrest in G2. .

38

Figure 1.7. NIMA kinase NIMA is a serine/threonine specific kinase which consists of an amino-terminal catalytic domain and a carboxy-terminal regulatory domain. The carboxy-terminus contains two potential coiled coil domains and three consensus nuclear localization signals. PEST sequences in this region are important for NIMA degradation as loss of these sequences produces a highly stabilized version of the kinase.

39 NIMA kinase activity

NIMA protein level

S G2 M G1

NIMA NIMA NIMA NIMA Inactive P P P P P P P Active Fully Proteolytic Active Destruction

(Courtesy Jon Davies, Adapted from Ye et. al, 1995)

Figure 1.8. Regulation of NIMA through the cell cycle NIMA protein levels and kinase activity fluctuate during the cell cycle. During interphase NIMA levels are very low and the kinase is inactive. Protein levels rise through G2 and reach a peak at mitosis. During G2 NIMA is phosphorylated and is activated. NIMA becomes hyper-phosphorylated and fully activated at the G2-M transition. During mitotic exit NIMA is rapidly degraded resulting in lowering protein levels and kinase activity.

40

(Reproduced with permission from O’Connell, et al. 2003)

Figure 1.9. The NIMA-related family of kinases NIMA-related kinases have been identified in a range of organisms from fungi through to humans. NIMA-related kinases have been identified largely on the basis of sequence homology between their kinase domains and the catalytic domain of NIMA. Many of the NIMA-related kinases contain regulatory domains in their carboxy-termini similar to NIMA, including coiled coil domains and PEST sequences.

41 S. pombe S. cerevisisae Locus Description ortholog ortholog

ankA (sntA) wee1 kinase,negative regulator of NimX Wee1 SWE1 bimA (sepI) Blocked in mitosis mutant,APC/C component 3 (APC3) nuc2 CDC27 bimB Blocked in mitosis mutant,separation of sister chromatids cut1 ESP1 bimC Blocked in mitosis mutant,BimC class kinesin cut7 KLP1/CIN8 bimD Blocked in mitosis mutant,DNA metabolism pds5 PDS5 bimE Blocked in mitosis mutant,APC/C component 1 (APC1) cut4 APC1 bimF Blocked in mitosis mutant bimG Blocked in mitosis mutant,phosphoprotein phosphatase dis2 GLC7 bimH APC/C component 6 (APC6) cut9 CDC16 bncA Binucleate conidia hfaB High frequency of aneuploids mipA c tubulin tug1 TUB4 nimA Never in mitosis mutant,serine/threonine protein kinase fin1 kin3 nimB Never in mitosis mutant nimC Never in mitosis mutant nimD Never in mitosis mutant nimE Never in mitosis mutant,cyclin B cdc13 CLB2 nimF Never in mitosis mutant nimG Never in mitosis mutant nimH Never in mitosis mutant nimI Never in mitosis mutant nimJ Never in mitosis mutant nimK Never in mitosis mutant nimL Never in mitosis mutant nimM Never in mitosis mutant nimN Never in mitosis mutant nimO Never in mitosis mutant,DNA Replication dfp1 DBF4 nimP Never in mitosis mutant nimQ Never in mitosis mutant,DNA Replication mcm2 MCM2 nimR Never in mitosis mutant nimS Never in mitosis mutant nimT Never in mitosis mutant,tyrosine phosphatase,positive regulator of NimX cdc25 MIH1 nimU Never in mitosis mutant nimV Never in mitosis mutant nimW Never in mitosis mutant nimX Never in mitosis mutant,protein kinase cdc2 CDC28 nuvF Mutagen sensitive,DNA synthesis checkpoint sldA Synthetic Lethal without Dynein,spindle checkpoint bub1 BUB1 sldB Synthetic Lethal without Dynein,spindle checkpoint bub3 BUB3 snoA Suppressors of nimO snoB Suppressors of nimO sntB Suppressors of nimT sntC Suppressors of nimT snxA Suppressor of nimX snxB Suppressor of nimX snxC Suppressor of nimX snxD Suppressor of nimX sonA Suppressor of nimA1,nucleocytoplasmic transport rae1 GLE2 sonB Suppressor of nimA1 sudA Suppressor of bimD6,chromosome sca .old protein psm3 SMC3 sudB Suppressor of bimD6 sudC Suppressor of bimD6 sudD Suppressor of bimD6,chromosomal condensation SPAC10F6.10 RIO1 tinA Two-hybrid interacting with NimA uvsB UV sensitive,DNA damage checkpoint rad3 MEC1 uvsD UV sensitive,DNA damage checkpoint rad26

(Adapted from Osmani and Mirabito 2004)

Table 1.1. List of A. nidulans cell cycle mutants

42

CHAPTER 2

MATERIALS AND METHODS

2.1. General DNA preparation and cloning 2.1.1. Plasmid maxiprep and miniprep Plasmid minipreps were initiated by inoculating a bacterial colony from a fresh plate into 2 ml 2xTYP (16 g/L yeast extract, 16 g/L tryptone, 5 g/L sodium chloride, 2.5 g/L potassium phosphate, 490 mg/ L magnesium sulfate, [pH 7.4]). Cultures were grown overnight at 37°C with 225rpm shaking in an air shaker (Innova). Minipreps were performed using a miniprep plasmid kit according to the manufacturers’ instructions (Promega). Plasmid maxipreps were initiated by inoculating 1 ml of an overnight starter culture into 100 to 1 l of Luria-Bertaini broth (LB) (10 g/L bacto tryptone, 5 g/L yeast extract, 5 g/L sodium chloride) or 2xTYP broth. Cultures were grown overnight at 37°C with 200 rpm shaking in an air shaker (Innova). Maxipreps were perfomed using a maxiprep plasmid kit according to the manufacturer’s instructions (Promega).

2.1.2. DNA cloning Restriction digests were conducted using commercially available endonucleases (New England Biolabs and Promega). Digests were performed according to the manufacturers’ protocols in supplied buffers. If desired enzyme sites were not present in the insert DNA, restriction sites were added through PCR by using primers possessing sequence homologous to the insert DNA as well as 5’ extensions containing desired restriction sites. Restriction digests were analyzed by agarose gel electrophoresis.

43 Restriction fragments were readied for ligation reactions either by agarose gel purification using a Gel Extraction Kit (Qiagen) or using a DNA Clean Kit (Qiagen). For ligation, vectors and inserts were digested with appropriate restriction endonucleases. Linearized vector and insert were combined using a 1:4 molar ratio in combination with 70 Units T4 DNA Ligase (New England Biolabs) and 1x T4 Ligase Buffer (New England Biolabs) in a final volume of 10 μl to 20 μl. Ligation reactions were incubated overnight at 16°C.

2.1.3. Polymerase Chain Reaction (PCR) PCR was performed using a 9700 Thermal Cycler (Perkin Elmer) or a Gradient Thermal Cycler (Eppendorf). PCR cycling conditions varied based on the protocol and are described in the individual methods descriptions. Generally, Pfu Turbo Polymerase (Stratagene) or Vent Polymerase were used for high fidelity PCR reactions for cloning. The Expand Long Template PCR Kit (Roche) or Taq DNA Polymerase (Sigma) was used for PCR screening protocols.

2.1.4. Primers Oligonucleotide primers for sequencing, tagging, and cloning were designed using the Primer Designer Ver. 2.0 software (Scientific and Educational Software). Primer sequences are listed in Table 2.1

2.1.5. DNA sequencing DNA sequencing was performed by the Plant Microbial Genomics Facility at The Ohio State University (Columbus, OH). 1µg of plasmid DNA was provided along with 10pmoles of each primer dilutes as 2pmol per µL. Analysis of the sequencing results were carried out with Sequencing Analysis software Ver 3.3 (Perkin Elmer Applied Biosystems) or DNAStar Seqman software (DNAStar).

44 2.1.6. Bacterial strains DH5αF’ competent Escherichia coli were used for all general cloning and plasmid amplification applications during suppressor plasmid re-isolation. Additional strains of bacteria (GC5 and genetech) were used in conjunction with various kits, and are identified in the manufacturers’ protocols.

2.1.7. Transformation of bacteria DH5αF’ competent E. coli cells were used for general plasmid amplification applications. Competent cells were stored in 100 μL aliquots at -80°C. Immediately prior to transformation, competent cells were thawed on ice. 20ng super-coiled plasmid DNA was added directly to the cells and mixed gently. When transforming plasmid DNA from low-melt agarose, the agarose was warmed at 65OC for 15 minutes and 30 μL TCM (10 mM Tris [pH 7.5], 10 mM Calcium chloride, and 10 mM magnesium chloride) was added prior to addition to competent cells. After addition of DNA, cells remained on ice for an additional 15 minutes and then were shifted to a 42OC water bath for 2 minutes. Following the heat shock, cells were returned to ice for an additional 2 minutes. 900 μl of 2xTYP was added to each tube and the cells were allowed to recover for 1 hour in a shaking air incubator at 37OC and 225 rpm. Transformed cells were plated onto LB agar plates containing appropriate antibiotic selection in 100 μL and 900 μL aliquots. The cells were incubated for at least 15 hours at 37OC in an air incubator.

2.1.8. Storage and stock preparation of bacteria 3 ml of 2xTYP with appropriate antibiotic were inoculated from a single bacterial colony using a sterile applicator. Cultures were grown for at least 16 hours at 37OC with shaking at 225 rpm in an air incubator. Cells were collected from media by centrifugation for 2 minutes at 14,000 rpm. The bacterial pellet was resuspended in 1 mL of 2xTYP and 20% glycerol in a sterile 1.5 mL Eppendorf tube and was stored indefinitely at -80OC.

45 2.2. Culture and genetics of A. nidulans 2.2.1. A. nidulans specific media YG media: (56 mM dextrose, 5 g/L yeast extract, 10 mM magnesium sulfate, supplemented with 1 µg/ml p-aminobenzoic acid (paba), 0.5 µg/ml pyrodoxine HCL (pyro), 2.5 µg/ml riboflavin HCL (ribo), 2 µg/ml nicotinic acid, 20 µg/ml choline, 20 ng/ml D-biotin and 1 ml/l trace elements). Strains carrying uncomplemented pryG89 auxotrphic mutation were grown in YGUU (YG media supplemented with 1.2 g/l uridine and 1.12 g/l uracil. YAG media: (YG media with 1.5% w/v agar) MAG media: (20 g/l malt extract, 20 g/l bacto peptone, 56 mM dextrose, supplemented with 1 µg/ml p-aminobenzoic acid (paba), 0.5 µg/ml pyrodoxine HCL (pyro), 2.5 µg/ml riboflavin HCL (ribo), 2 µg/ml nicotinic acid , 20 µg/ml choline, 20 ng/ml D-biotin, 50 mg/l adenine sulfate, 50 mg/l leucine, 50 mg/l L-methionine, 100 mg/l arginine, 200 mg/l L-lysine HCL, 1 ml/l trace elements and 2% agar). Strains carrying uncomplemented pryG89 auxotrphic mutation were grown on MAGUU (MAG media supplemented with 1.2 g/l uridine and 1.12 g/l uracil.) Minimal Media Urea: (10 mM urea, 7 mM potassium chloride, 1 mM magnesium sulfate, 1 ml/l trace elements, and supplements as required). Glucose (final concentration 1% w/v) or glycerol (final concentaration 0.47 % v/v) was added prior to autoclaving. Ethanol (final concentration 1% v/v) was added after autoclaving. After autoclaving add potassium phosphate [pH 6.8] to 12 mM and sodium thiosulfate to 3.2 mM. For solid media 1.5% w/v agar was added prior to autoclaving. Minimal media Low Nitrate: (82 mM sodium nitrate, 7 mM potassium chloride, 2 mM magnesium sulfate, 11 mM potasssium phosphate monobasic, 111 mM dextrose, 1 ml/l Clive Roberts Trace Elements, additional supplements as required, and 1.5% w/v agar [pH 6.7]). Minimal media Yeast Lactose: (10 mM urea, 7 mM potassium chloride, 1 mM magnesium sulfate, 5 g/l yeast extract, 20 g/l lactose, 1 ml/l trace elements, and supplements as required) After autoclaving add potassium phosphate [pH 6.8] to 12 mM

46 and sodium thiosulfate to 3.2 mM. 40 mM threonine was included for alcA::based protein induction.

2.2.2. Preparation of A. nidulans conidia stock suspensions Haploid A. nidulans strains were inoculated at a concentration of 1X107 spores/ml into four mls of MAG media (strains carrying un-complemented pyrG89 were supplemented with uracil and uridine) containing 0.9% agar at 48°C. Inoculated media was vortexed to mix the suspension and overlayed onto MAG plates. Innoculated plates were incubated at 30°C for no more than 40 hours to allow for conidiation. Conidia were harvested from the surface of the plates in 10 ml of 0.2% Tween 20. A sterile glass spreader was used to gently rub the top of the fungal lawn to release conidia into suspension. Suspended conidia were transferred to sterile 15 ml polypropylene tubes (Corning). Conidia suspensions were at centrifuged at 7,000 rpm for 2 min. to pellet

conidia. Conidia were washed three times in 15 ml of dH2O. After the final wash conidia were separated from hyphal debris by gently resuspending only the top conidial layer of the pellet in stock storage solution (8.5 mM sodium chloride, 200 µM Tween 80 (Fisher Scientific). Conidial suspensions could be stored for a month at 4°C.

2.2.3. Conidiospore Quantitation Conidial suspensions of A. nidulans were quantitated to allow for accurate inoculation densities for germination in liquid media. Conidial suspensions were quantiated by counting 10 μL of a 1 x 10-3 dilution of conidia in 0.2% Tween 20 using a Bright-Line hemocytometer (Reichert-Jung). Four fields of conidia were counted for each sample and the average value used for quantitation. The number of conidia obtained from this count was multiplied by 1 x 107 conidia per ml to determine the concentration of the original suspension.

47 2.2.4. Long term Storage and Stock Preparation of A. nidulans Strains were streaked to single colonies on selective three times. Conidia from these colonies were harvested using a sterile wire loop, which had been immersed in 0.2% Tween, and streaked out on selective media. Inoculated agar cultures were incubated at 37OC for 48 hours and then at room temperature for 3 days. 1 mL of sterile

7.5% milk (7.5g of Carnation Nonfat Dry Milk in 100mLs dH2O and autoclaved for 20 minutes) was applied to the surface of the cultures, and harvesting of mature conidia was accomplished by agitation of the colony surfaces using a sterile glass spreader. A sterile transfer pipette was used to transfer 250 μL of the suspended spores into two screw top vials containing baked silica. The silica was vortexed briefly to assist in even distribution of the spores and returned to ice for 1 hour. The silica was left at room temperature for 2 days with the vial lid loosened to allow for complete drying. Both vials were placed in a room temperature desiccator. Strains were regrown from silica stocks by pouring several silica pieces onto appropriate agar, and incubating at 32°C for a minimum of 48 hours in an air incubator.

2.2.5. Strain generation by meiotic crossing A. nidulans strains were induced to undergo meiosis by ensuring that each strain contained at least one forcing auxotrophic marker which was complemented in the other strain. The use of parental strains with differing conidial colors provided a visual screen for successful meiotic crossing events. Parental strains were alternately spot inoculated onto MAGUU media with approximately 2 cm between spot inoculations. Inoculated plates were incubated at 30°C until the edges of adjoining colonies abutted each other. A strip of hyphae at the interface of the two colonies was removed using sterilized tweezers. This hyphal mat was crushed onto the surface of a minimal media low nitrate plate. The plate was sealed with tape and incubated at 30°C for a minimum of 2 weeks to allow cleistothecia to form. Cleistothecial maturation was monitored using a dissecting microscope (Bausch and Lomb). Mature cleistothecia were removed with a sterilized glass pipette and rolled across the surface of a 4% water agar plate to clean any debris from the surface of the 48 cleistothecia. Once cleaned, cleistothecia were crushed in 0.2% Tween 20 in a 1.5 ml Eppendorf tube to release ascospores. Ascospores were plated on MAGUU to determine whether the strains had crossed. Individual colonies were selected and tested on a range of minimal media plates lacking various supplements to identify strains with desired genotypes.

2.3. General A. nidulans techniques 2.3.1. Small scale protein preparation Conidia were inoculated at roughly 1x106 conidia/ml concentration into 30 mL of YG or minimal media in sterile petri dishes. Cultures were incubated overnight at 30°C until just before hyphae began to conidiate at the media air interface. Mycelia were harvested through Miracloth as described for large scale protein preparation. Mycelia were frozen in liquid nitrogen and dried overnight in a lyophilizer (Savant). The next day samples were pulverized with toothpicks. The mycelia were weighed and mixed with 2X 6M Urea Sample buffer (250mM Trizma Base [pH6.8], 7 M Urea, 100 µl/ml β- mercaptoethanol, 200 µl/ml glycerol, 4% sodium dodecyl sulfate) at 40 μl/mg of dried mycelia. Samples were boiled for 5 min. prior to analysis by SDS-PAGE.

2.3.2. Large scale protein preparation Conidia were inoculated into appropriate liquid media at 2x106 conidia/ml concetration. Culture flasks were coated with GelSlick (Cambrex Bio Science) prior to culture addition to prevent mycelia from adhering to the sides of the flask. Cultures were frequently germinated overnight at 28°C with agitation in an air incubator. The next day the temperature was increased to 32°C and cultures were grown to early log phase. Culture growth was monitored by performing packed cell volume measurements. 10 ml of culture was removed and centrifuged for 1 min. at 3200 rpm in a clinical swinging bucket centrifuge (Thermo IEC). Packed cell volumes of between 0.2 to 0.3ml of mycelia indicated that the culture was in log phase growth. Mycelium was harvested by vacuum filtration of the culture through Miracloth (Calbiochem). The mycelium was washed twice with Stop Buffer (9 g/L NaCl, 65 mg/L NaN3, 20 mL 0.5 M EDTA [pH

49 8.0], 2.1 g/L NaF). Excess liquid was pressed out of mycelia and the mycelium was removed from the surface of the Miracloth. Harvested mycelium was placed in a sterile polypropylene tube (Corning) and immersed in liquid nitrogen and dried overnight in a lyophilizer (Savant). The next day samples were pulverized by grinding the dried mycelia with a mortar and pestle and 1.5 mL of HK buffer (1 µl/ml leupeptin (10 mg/ml in dH2O), 0.4 µl/ml soybean derived trypsin and chymotrypsin inhibitor (25 mg/ml in DMSO), 0.5 µl/ml N-tosyl-L-phenylalanine chloromethyl ketone (TPCK) (50 mg/ml in DMSO), 5 µl/ml Aprotinin (1.5mg/ml), 760 µg/ml N-p-tosyl-arginine methyl ester hydrochloride (TAME), 6.2 mg/ml p-nitrophenyl phosphate (PNPP), 800 µg/ml benzamidine, 200 µg/ml sodium vanadate, 420 µg/ml sodium fluoride, 71µl/ml 1 M β- glycero-phosphate, 35 µl/mL 0.5 M Ethylene-glyco-tetra-acetic acid (EGTA), [pH 8.0], 12 µl/mL 0.5 M ethylene-diamine-tetra-acetic acid (EDTA) [pH 8.0], 30 µl /ml 1.0 M Tris-HCL, [pH 7.5], 24 µl/ml 10% Nonidet P-40.) /100 mgs of mycelia and vortexed. The mixed mycelial slurries were aliquotted into 50 ml centrifuge tubes and centrifuged at 21,000 rpm at 4°C for 30 min to pellet mycelial debris. The supernatant was transferred to fresh falcon tubes and protein estimation done using standard Bradford

method using Bio-rad protein assay reagent and by noting absorbance at O.D595. Protein samples were further used for affinity purification purposes and finally eluted with SDS sample buffer (625mM Tris-Cl pH 6.8, 4% SDS, 20% glycerol, 10% beta- mercaptoethanol).

2.3.3. Small scale genomic DNA extraction A small amount of spores were inoculated in petri plates with appropriate media and the grown mycelia harvested with Stop Buffer (9 g/L sodium chloride, 65 mg/L sodium azide, 20 mL 0.5 M EDTA [pH 8.0], 2.1 g/L sodium fluoride). Excess liquid was pressed out of mycelia and the mycelia was removed from the filter, placed in an eppendorf and immediately immersed in liquid nitrogen. The mycelia was removed from the liquid nitrogen and lyophilized overnight. The dried mycelia were pulverized and 100 µl of Miniprep Lysis Solution (Promega) was added and vortexed to mix. 100 µl Miniprep Neutralization Solution was added and vortexed to mix. Samples were

50 centrifuged at 14,000 rpm in a Model 5420 table top refrigerated centrifuge (Eppendorf) at 4°C for 10 minutes. The supernatant was processed using Miniprep Purification Kit (Promega) according to the manufacturer’s instructions. Genomic DNA was eluted from the column in 50 µl dH2O.

2.3.4. Large scale genomic DNA extraction A. nidulans conidia were inoculated into appropriate media and incubated overnight in an air incubator at 32OC at 180 rpm. Inocula were allowed to grow until a 10 mL sample yielded a packed cell volume of 0.5 mL after undergoing centrifugation at 7,000 rpm. The mycelium was harvested by vacuum filtration of the media through Miracloth (Calbiochem). The mycelium was washed twice with Stop Buffer (9 g/L sodium chloride, 65 mg/L sodium azide, 20 mL 0.5 M EDTA [pH 8.0], 2.1 g/L sodium fluoride). Excess liquid was pressed out of mycelia and the mycelium was removed from the filter, placed in a 50 mL polypropylene tube (Corning) and immediately immersed in liquid nitrogen for at least 2 minutes. The mycelium was removed from the liquid nitrogen and lyophilized overnight. Lyophilized mycelia samples were stored at -80OC until DNA extraction. To extract genomic DNA, 20 mg of lyophilized mycelia was thoroughly ground with a disposable pestle in a 1.5 mL Eppendorf microcentrifuge tube. After grinding, 250 μL 0.5% SDS DNA Extraction Buffer (200mM Trizma Base [pH 8.5], 250 mM NaCl, 25 mM EDTA, 0.5% SDS), 175μL phenol, and 75 μL chloroform were added directly to 40 mg of ground mycelia. Tubes were rocked for 15 minutes at room temperature, after which, debris was pelleted by centrifugation at 14,000 rpm for 20 minutes. 400 μl of chloroform was added to the supernatant and spun at 14,000 rpm for 10 minutes. The supernatant was combined with an equal volume of 5 M lithium chloride, placed on ice for 10 minutes to precipitate RNA, and centrifuged at 14,000 rpm for 10 minutes. DNA was precipitated from the supernatant using 2-propanol and resuspended in 50 μL TE.

51 2.3.5. Transformation of A. nidulans 1 x 109 fresh conidia were inoculated into 50 mL YGUU (YG media augmented with 1.2 g/L uridine and 1.12 g/L uracil). Liquid cultures were grown at 32OC at 200 rpm in an air incubator for 5.5 hours or until conidia had just begun to germinate. Germlings were harvested by centrifugation in a swinging bucket rotor at 2,000 rpm for 2 minutes and resuspended in a protoplasting mix containing 20 mL Solution 1 (105.6 g/L ammonium sulfate, 19.2 g/L citric acid, [pH 6.0]), 20 mL Solution 2 (10 g/L yeast extract, 20 g/L sucrose, 1 µg/mL acid paba, 0.5 µg/mL pyro, 2.5 µg/mL ribo, trace

elements, 4.92 g/L MgSO4), 80 mg bovine serum albumin (BSA), 20 mg novozyme, 100 mg driselase and 100 μL β-glucoronidase. More recently, 10 mg/ml of the enzyme VINOFLOW FC was used in place of driselase and β-glucoronidase. The resuspended germlings were transferred to a clean sterile flask and incubated in an air incubator at 32OC and 170 rpm for 2 to 3 hours or until the cell wall has degraded to such an extent that large vacuoles, within released protoplasts, become visible under examination using light microscopy. Protoplasts were collected by centrifugation in a swinging bucket rotor for 2 minutes at 7,000 rpm. Protoplasts were washed two times in Solution 3 (52.8 g/L ammonium sulfate, 10 g/L sucrose, 9.6 g/L citric acid, [pH 6.0]) and resuspended in 1 mL

Solution 5 (44.7 g/L KCl, 7.35 g/L CaCl2, 2.09 g/L MOPS, [pH 6.0]). Transformation was carried out by combining 2-4 μg DNA, 100μL protoplasts,

and 50 μL of room temperature Solution 4 (250 g/L PEG 8000, 7.35 g/L CaCl2, 44.7 g/L KCl, 10 mL 1 M Tris [pH 7.5]). The transformation reaction was incubated on ice for 20 minutes before addition of an additional 1 mL of Solution 4. After an additional 20 minutes of room temperature incubation, the transformations were plated onto YAG sucrose (5 g/L yeast extract, 3.6 g/L dextrose, 342.3 g/L sucrose, 2.47 g/L MgSO4, 1μg/mL PABA, 500 ng/mL pyro, 2.5 μg/ml ribo, trace elements, 15 g/L agar) in 4 mL YAG sucrose overlays (same as above except 10 g/L agar) in 10 μL, 50 μL, 100 μL, 250 μL, and 500 μL volumes. Transformation plates were incubated in an air incubator at 32°C for 70 hours or until colonies had developed.

52 2.3.6. Immunofluoresence A. nidulans strains were inoculated into liquid media at concentrations of 1x105 to 1x106 conidia / ml depending on the strain being examined. Inoculation densities were higher in minimal media than in YG media. 300 µL inoculated media was pipetted onto No. 1 ½ coverslips (Corning Labware & Equipment) inside 100x15mm petri dishes (Fisher Scientific). Up to twelve coverslips were placed in each petri dish. Strains were allowed to germinate at 25°C to 37°C, depending on the strain and media conditions untl germlings had reached the desired length. Samples were fixed by inversion of the coverslip on 150 µl of immunofluorescence fix (6% paraformaldehyde, 0.1% glutaraldehyde, 5%DMSO, volume up to 25.5 ml with PHEM (45mM PIPES, 45mM HEPES, 10mM EGTA, 5mM magnesium chloride, [pH 6.9])) for 45 min at room temperature. Coverslips were washed thee times in a volume of at least 150 mL PHEM. Digestion of A. nidulans cell wall components was accomplished by inversion of coverslips on 150 µL of immunofluorescence mix (8 mg/ml β-D-glucanase (Interspex Products, Inc.) 1 mg/ml Liticase (ICN Biomedicals, Inc.), 5 mg/ml Driselase (InterSpex Products, Inc.), 50 mM sodium citrate [pH 6.0], 1 mM magnesium sulfate, 2.5 mM EGTA, and 2% BSA) at 30°C. Digestion was monitored by removal of samples at 10 min intervals following digestion for 1.5 hours. For cells germinated in YG media 2X immunofluorescence digest mix was used. Cover slips were washed three times in PHEM. Primary antibody incubations were carried out by inversion of cover slips in 150 µL of immunofluorescence buffer (PHEM, 0.01% Tween20, and 3% BSA) plus primary antibodies for one hour at room temperature. Primary antibody used in this study for visualization of the nucleolus included the freeze dried autoantibody control ANA positive human serum (Sigma Diagnostics) resuspended in 4000 µL PHEM + 3% BSA + 0.1% tween20 + Azide. No further dilutions were required and 200 µL of the above was used per coverslip. Cover slips were washed three times in PHEM. Secondary antibody incubations were carried out as described for primary antibodies. Secondary antibody used in this studies included Alexa Flour 594 goat anti-human (Molecular Probes, Inc.). Following secondary antibody incubation, cover slips were washed three times in PHEM and mounted in 11 µL of an 80% Citifluor solution with 150ng/µL DAPI. 53 2.4. Isolation of full-length cDNAs by RACE PCR To verify full-length cDNA clones for mcnA, mcnB and mcnC we used 5’ RACE PCR. To generate an A. nidulans cDNA library, A. nidulans total RNA was prepared from 2g lyophilized R153 mycelia using Ultraspec RNA Isolation System (BIOTECX). Lyophilized mycelia were ground using a ceramic mortar and pestle which was pretreated

in 2 % H2O2 for 1 hour to eliminate RNAse activity. Ground mycelia was combined with 40 mL of Ultraspec RNA reagent and placed on ice for 5 minutes. 8 mL chloroform was added and the mycelia were placed on ice for an additional 5 minutes. Mycelia were centrifuged at 14,000 rpm for 15 minutes to remove debris. RNA was precipitated using

2-propanol and resuspended in 1.5 mL diethyl pyrocarbonate (DEPC) treated H2O. The total RNA concentration (24 μg/μL) was determined by measuring the absorbance of a

sample of total RNA at O.D.260. Poly-A+ mRNA was isolated using the PolyATtract mRNA Isolation System

(Promega). 9.6 mg of total RNA was added to 4.46 mL of DEPC H2O and placed in a 65O C waterbath for 10 minutes. 10 μL of Biotinylated-Oligo (dT) Probe and 60 μL of 20xSSC were added to the RNA, and incubated at room temperature until cool. The entire annealing reaction was incubated with paramagnetic particles and mRNA was

isolated by exposing particles to a magnet. mRNA was eluted in 500 μL of DEPC H2O at a final concentration of 136 μg/mL. Therefore, isolated mRNA represented 0.7% of the total RNA. This is consistent with previous findings for A. nidulans. cDNA synthesis was performed using 1 μg Poly-A+ mRNA and cDNA synthesis reagents from the Marathon cDNA Amplification kit (Clontech) Full-length mcnA, mcnB and mcnC cDNAs were isolated by 5’ RACE PCR using the Touchdown PCR Method as described in the Marathon cDNA Amplification protocol (Clonetech). PCR was performed using an AP1 adaptor ligated cDNA library, an adapter primer (complementary to the adaptor sequence), and gene specific primers for all three genes. The full-length cDNAs were cloned into the PCR 2 or PCR 2.1 vectors (Invitrogen) using the TA Cloning System (Invitrogen). cDNAs were completely sequenced.

54 2.5. Fusion PCR The fusion PCR-based method of amplifying gene-targeting constructs solves the problem of eliminating the need to perform ligation reactions required in conventional cloning strategies. This method employs to produce two different types of gene-targeting constructs used to generate A. nidulans strains.

2.5.1. Gene deletion constructs To delete the entire open-reading frame of a given gene, initially three different DNA fragments were amplified. The first fragment is the gene deletion cassette consisting of the heterologous (but functionally complementary) A. fumigatus pyroA gene (Af-pyroA) to minimize the chance of the eventual targeting construct integrating into the native A. nidulans pyroA locus upon transformation. This fragment was amplified using appropriate primers listed in table 2.1 as ΔP1 and ΔP2 with the plasmid pUC19-pyroA. In addition, two fragments of the genome sequences immediately upstream and downstream of the gene (~1.0KB in length) to be deleted were PCR amplified as targeting regions using appropriate primer pairs listed in table 2.1 as 5’fP3/5’rP4 and 3’fP5/3’rfP6. In amplifying these two fragments, the 3’-end primer of the upstream sequence is designed to have a 20 bp long 5’ extension that is a reverse complement of the first 20 bp sequence of the Af-pyroA cassette (Figure 2.1, shown in red). The 5’-end primer of the downstream sequence is designed to have a 20 bp long 5’ extension identical to the last 20 bp sequence of the Af-pyroA cassette. A fusion PCR reaction using a mixture of these three fragments (A, B and C) in equal molar amounts (estimation deduced by DNA concentration and length of the fragments) as templates and the 5’-forward primer (5’fP3) of the upstream sequence and the 3’-reverse primer (3’rP6) of the downstream sequence was used to amplify the complete gene deletion construct with the three initial fragments fused into a single DNA fragment (Figure 2.1)(Yang, Ukil et al. 2004). The PCR mix was a total volume of 50μL, including 0.75μL Expand Long Template enzyme mix, each dNTPs at 500μM final concentration, 20ng pFN03 plasmid template DNA or 100ng of genomic DNA, 5μL 10X buffer 3, each 300nM (final concentration) of the primers FN02 and LU212. The PCR cycling conditions were as

55 following. 94°C 2 minutes, 25 cycles (94°C 15 seconds, 55°C 30 seconds, 68°C 2 minutes), 68°C 5 minutes. The gene specific primers used to amplify the 5’-flanking as well as the 3’-flanking sequences of all phosphatase genes are listed in Table 2.1. The basic PCR cycling conditions for amplifying these fragments were as following with minor changes introduced for individual genes. 94°C 2 minutes, 25 cycles (94°C 15 seconds, 55°C 30 seconds, 68°C 2 minutes), 68°C 5 minutes. Once all three fragments for fusion PCR were generated, 500ng of each of the three fragments were used a template DNA for fusion PCR reaction. The PCR mix was a total volume of 50μL, including 0.75μL Expand Long Template enzyme mix, each dNTPs at 500μM final concentration, 500ng of each template DNA, 5μL 10X buffer 3, each 300nM (final concentration) of the appropriate primers (for conceptual layout of fusion PCR see Chapter 3 Introduction). The basic PCR cycling conditions for amplifying these fragments were as following with minor changes introduced for individual constructs. 94°C 2 minutes, 25 cycles (94°C 15 seconds, 55°C 30 seconds, 68°C 6 minutes + 15 seconds increment each cycle), 68°C 5 minutes. Upon transformation of this construct, transformant colonies are tested for proper replacement by making small scale genomic preparation and performing a diagnostic PCR using appropriate primer pairs listed in table 2.1 as P7 and P8 that targets sequences outside of this construct in diagnostic PCR reactions.

2.5.2. Endogenous C-terminal tagging constructs Generation of endogenous C-terminal tagging constructs by fusion PCR is identical to the generation of deletion constructs except that it involved the insertion of a sequence encoding the tag instead of deletion of a gene. In order to insert a GFP or S tag at the C-terminus of a protein of interest, the two targeting fragments amplified for homologous recombination are ~1.0KB in sequence and encode the genomic sequence upstream of the termination codon and the immediate downstream genome sequence following the termination codon of the gene. In amplifying these targeting fragments, the 3’-end primer of the upstream sequence is designed to have a 20 bp long 5’ extension that is a reverse complement of the first 20 bp sequence of the tagging cassette, and the 5’-end

56 primer of the downstream sequence is designed to have a 20 bp long 5’ extension identical to the last 20 bp sequence of the tagging cassette. The tagging cassette itself is made up of the sequences encoding a GA5 (five-time repeat of glycine-alanine residues) linker immediately followed by a sequence encoding a GFP polypeptide and an Af-pyrG auxotrophic marker. The GA5 linker is a flexible bridge translationally fusing the protein of interest and the GFP polypeptide, minimizing the interference of the functional conformation of the two protein entities (Osmani et al., 2006a). Both the GFP tagging cassette as well as the affinity S-tagging cassette were amplified with the approrpriate primers listed in table 2.2 named as either GFP-P1/GFP-P2 or S-P1/S-P2 using plasmid pFNO3 (for GFP) and plasmid pAO81 (for S-tag) (In Figure 2.2, the primers are represented as D and E). The gene specific primers used to amplify the 5’-flanking (termed 5’ GFP-fP3/rP4 and 5’ S-fP3/rP4) as well as the 3’-flanking (termed 3’ GFP- fP5/rP6 and 3’ S-fP5/rP6) sequences of all genes whose encoded proteins were tagged are listed in table 2.2. PCR conditions were similar as in generating the deletion constructs using external primers P3 and P4 (Figure 2.2, represented as F). The final fusion PCR reaction is carried out using a mixture of the three fragments as templates and the 5’-end primer of the upstream sequence and the 3’-end primer of the downstream sequence with conditions similar to section 2.4. Once the tagging constructs were amplified, transformation was carried out according to the standard protocol using diagnostic primers termed GFP-P7/P8 or S-P7/P8 (Figur 2.2, represented as G). Note for the gene CgrA alone, tagging was done using the pJH19 plasmid to make a ref fluorescently tagged protein.

2.6. S-tag purification in A. nidulans After large scale purification of proteins, 300µl of S-protein Agarose slurry (150ul packed beads volume) [Novagen # 69704] was added per 100mg of total protein. The solutions were incubated on ice at 4°C for 2 hours while gently rocking. After incubation, the S-protein beads were recovered by centrifugation at 3,800rpm for 2 minutes in a clinical swinging bucket centrifuge (Thermo IEC). The spun-down beads were washed with equal volume of protein wash buffer 5 times (i.e., if total sample

57 volume of S-tag pull down solution was 10ml, it was washed with 10ml of wash buffer. The protein wash buffer was identical to the protein extraction buffer icluding NaCl [final concentration 300mM]. After washing, the beads were transferred to 1.5mL Eppendorf tubes and washed at least 2 times more, all at 4°C. 4X sample buffer were added and the samples were boiled for 5 minutes. The samples were vortexed and the beads removed from the supernatant by centrifugation for 5 minutes at 14,000rpm. The supernatants were taken and loaded onto 10% polyacrylamide protein gels.

2.7. Coomassie staining of protein gels After gel electrophoresis was complete, polyacrylamide protein gels were fixed in 50% ethanol and 10% acetic acid overnight. The next day, the fixed gels were rinsed 3

times for 5 minutes with dH2O. Residual water was removed as much as possible, and Biorad Bio-Safe Coomassie solution (enough amount of to cover the gel) was added. Staining was continued for 1 hour, and background staining was removed by destaining in water for 30 minutes.

2.8. Silver staining of protein gels After gel electrophoresis was complete, polyacrylamide protein gels were fixed in 50% ethanol and 10% acetic acid for 10 minutes. The fixed gels were washed in 50%

ethanol for 15 minutes, and subsequently washed in dH2O three times for 5 minutes each.

After washing, the gels were incubated in 0.02g/100mL of Na2S2O3·5H2O solution for 1 minute. This was followed by washing in dH2O for three times 30 seconds each. Then the

gels were incubated in a silver nitrate solution (AgNO3 2g/L +750μl 37% formaldehyde

solution) for 20 minutes. The gels were washed in dH2O twice for 20 seconds each, and

after that the bands were visualized in the developing solution (Na2CO3 6g/100mL, 37%

formaldehyde 50μL/100mL, Na2S2O3·5H2O (0.02g/100mL) 2mL/100mL). When bands became visible the developing reaction was stopped with 50% ethanol and 10% acetic acid.

58 2.9. In vitro λ-phosphatase assay After purification of proteins via S-tag affinity purification, three tubes with equal amount of proteins (100mgs total protein) were prepared for no phosphatase, phosphatase and phosphatase + phosphatase inhibitors treatment. To initiate the λ-phosphatase assay (New England Biolab), the individual samples were washed in 2X phosphatase buffer

(i.e. volume of phosphatase buffer equal to volume of beads) which included MnCl2 and proper protease inhibitors (prepared according to manufacturer’s protocol). This step served the purpose of equilibrating the S-protein beads in the phosphatase buffer and removing phosphatase inhibitors present in the protein extraction buffer. After washing, the three samples were treated with either control buffer only or equal volumes of 1x phosphatase mix; no phosphatase, phosphatase (40 units λ-phosphatase), phosphatase (40 units λ-phosphatase) and phosphatase inhibitors (1mM Na Vanadate 50mM Na Fluoride). The beads were mixed with the solutions and incubated at 30°C for 30 minutes (mixing occasionally). Once the treatment was completed, sample buffer was added to a final concentration of 1X and boiled to stop the reaction. The supernatant was collected after pelleting of the beads by centrifugation at 14,000rpm and analyzed by SDS-PAGE.

2.10. alcA driven protein expression in A. nidulans Genomic DNA fragments were PCR amplified and cloned into alcA expression vectors pAL5 and pAL3 respectively as Kpn1/BamH1 restriction fragments. Constructs were transformed into A. nidulans by complementation of the pyrG89 auxotrophic marker. Transformants were screened for inhibition of colony formation as follows. Transformants, randomly selected were spotted onto YAG plates and incubated at 32°C to allow colony formation. Colonies were replica plated onto repressing (minimal media with 1% glucose) and inducing media (minimal media with 1% ethanol). Plates were incubated at 32°C to allow for colony formation, and growth inhibition on inducing media was compared to empty vector control strains. For immunofluorescence, transformants were germinated in either minimal media glucose to block protein expression, minimal ethanol to examine overexpression defects,

59 or minimal media including glycerol (4.66 ml/l), a non-inducing, non-repressing carbon source. For induction of alcA driven protein expression in large liquid cultures for protein preparation, cells were grown in minimal media yeast extract lactose media supplemented with 40mM threonine.

2.11. Measuring NIMA1 levels in nimA1 suppressed strains nimA1 strain (LPW3) transformed with AMA1 plasmid containing mcnA, mcnB, mcnC, wild type nimA or no insert (empty vector) were grown in YAG media at 32C till a packed cell volume of 0.4 was attained. At this point they were transferred to 42C, the restrictive condition for nimA1 mutation. To arrest nimA1 suppressed cells in mitosis in order to measure NIMA1 protein levels at their peak, 100g/mL of the drug Nocodazole was added after 15 minutes resulting in the activation of the SAC. The mycelia were harvested after 1 hour harvested by vacuum filtration of the media through Miracloth (Calbiochem). The mycelium was washed twice with Stop Buffer (9 g/L sodium chloride, 65 mg/L sodium azide, 20 mL 0.5 M EDTA [pH 8.0], 2.1 g/L sodium fluoride). Excess liquid was pressed out of mycelia and the mycelia was removed from the filter, placed in a 50 mL polypropylene tube (Corning) and immediately immersed in liquid nitrogen for at least 2 minutes. The mycelia was removed from the liquid nitrogen and lyophilized overnight and processed the next day for small scale protein preparation for western blot analysis. The primary antibody used was polyclonal NIMA antibody (E15III), 1:4000 dilutions. The secondary antibody used was 1:5000 of ECL peroxidase labeled anti-rabbit antibody (GE healthcare). Anti-tubulin antibody (B512), 1:5000 dilutions was used to analyze the same blot to assess loading control. The secondary antibody used was 1:5000 of ECL peroxidase labeled anti-mouse antibody (GE healthcare).

2.12. Western blot analysis Western blot analysis was performed to estimate protein expression levels and to verify proper integration of targeting constructs by confirming the protein size of chimeric protein. Small scale or large scale prepared proteins in appropriate sample

60 buffer were loaded on 7-10% SDS-polyacrylamide gels along with rainbow recombinant protein molecular weight marker (GE Healthcare). Gel electrophoresis was done at 30mAmp current and proteins subsequently transferred to nitrocellulose membrane using protein gel transfer apparatus at 180mAmp current for 3 hours. The membrane was further processed for protein detection by blocking in 5% milk (5.0 g Carnation Nonfat Dry Milk in 100mL of TBS buffer – 200mM Tris base, 5M NaCl pH 7.5) for 1 hour followed by 1 hour incubation with primary antibody made in 5% milk. The blot was then washed 3X TBST buffer (TBS buffer with 0.5% Tween 20). Secondary antibody (in 5% milk) incubation was performed for one hour followed by 3X washes in TBST buffer. The enhanced chemiluminescence reagent kit for western blot analysis was used to finally detect protein signal. The primary antibodies used were anti-NIMA (E15III – 1:5000 dilution), anti- GFP (Living colors – 1:5000), anti-DsRed (Living colors - 1:10,000), anti-S (Immunology consultancy laboratory – 1:5000) and anti-tubulin (Sigma B512 – 1:5000). Secondary antibodies respectively used were anti-rabbit (1:5000), anti- mouse (1:5000), anti-rabbit (1:5000), and anti- and anti-rabbit (1:5000). All secondary antibodies are ECL peroxidase labeled antibody from GE healthcare.

2.13. Site-directed mutagenesis In vitro site-directed mutagenesis was performed to generate the mutant MCNA- 3A. The DNA sequence to be mutagenised was amplified from the GFP tagged mcnA strain (LU256) using primers LU. The amplified DNA contained sequences for the mcnA gene fused to GFP and also incorporated the sequence for pyrGAF auxotrophic marker. The DNA was then cloned into the pCR-Blunt II-TOPO vector using the Zero blunt TOPO PCR cloning kit (Invitrogen). The vector containing the insert was then subjected to three rounds of site-directed mutagenesis following instructions in the Quickchange site-directed mutagenesis kit (Stratagene, catalog #200518) and using primer pairs – to convert serine 330, 465 to alanines. The mutations by site-directed mutagenesis were confirmed by sending the plasmid for sequencing. After confirmation of triple mutations, the insert was cleaved using RE BamHI and NotI and gel purified using the Qiagen gel purification kit. 1µg of insert DNA was then used to transform the mcnA deleted with

61 pyroA marker strain LU217 and by plating them on MAG plates with 1M sucrose. The transformant were further confirmed for proper landing of the insert at the mcnA locus by spotting them on MM + pyroA plates. Colonies that were pyrG plus but pyroA minus were considered positives. Refer section 6.2.5 for more details.

2.14. ΔmcnA, ΔmcnB and ΔmcnC phenotype testing The respective strains were tested for heterokaryon formation by replica streaking them on plates with and without selective pressure for the deletion marker. They were also tested for growth defects along with control strains R153 and JD100 under a variety of conditions. Strains were tested for their ability to undergo both self-crosses and crosses to other strains. Additionally strains were spot inoculated onto YAGUU media and tested phenotypes under the following conditions:1) temperatures of 20°C, 32°C, 37°C, and 42°C, 2) osmotic stress with 1M sucrose or 1M sodium chloride, 3) and chemicals including nocodazole (0.1 µg/ml, 0.2 µg/ml, 0.3 µg/ml, 0.4 µg/ml and 0.6 µg/ml), MMS (0.1% and 0.2%), and HU (4M, 6M, and 10M). All growth plates were incubated at 32°C except where indicated. These strains were also examined for cell cycle defects by DAPI staining. These strains were initially grown in either YGUU at 32°C.

2.15. Crosses between ΔmcnC and cell cycle mutants To determine whether loss of mcnC function was synthetically lethal with mutations in cell cycle regulatory genes, a ΔmcnC strain was crossed to strains carrying the following mutations: nimA1, nimA5. The double mutant strains were tested over a range of temperatures for synthetic lethality (20°C, 30°C, 32°C, 35°C, 37°C, and 42°C).

62 2.16. Examination of polarization defects in ΔmcnC expressing cells Strains containing mcnC deletion were inoculated into minimal media glucose at 5x105 conidia / ml. 300 µl of inoculum was placed on the surface of sterile cover slips and incubated at 32°C to allow germination. Cells were fixed and DAPI stained as described previously. To quantify the polarization defect, DAPI staining was used to count the number of nuclei and the length of the germtube measured by calibration on the eye piece. These were then compared to control cells from strain JD100. 100 cells were characterized for each strain.

2.17. Microscopy and image capture software Fixed samples were examined using an E800 microscope (Nikon, Inc.) with DAPI, FITC, and Texas Red filters (Omega Optical, Inc.). Image capture was performed using an UltraPix digital camera (Life Science Resources, Ltd.). To examine fluorescently tagged proteins in living cells, strains were germinated in 3 ml of minimal media lacking ribo in 35 mm glass bottom Petri dishes (MatTek Cultureware). Visualization of GFP/chRed fusion proteins was performed using a Nikon Eclipse TE300 (Nikon, Inc.) inverted microscope in conjunction with an Ultraview spinning disk confocal system (Perkin Elmer) and an Orca ER digital camera (Hamamatsu). Image capture on both microscopes was performed using Ultraview image capture software (Perkin Elmer).

2.18. Bioinformatics and DNA analysis Restriction analysis, DNA translation and identification of open reading frames was performed using Gene Runner Version 3.05 (Hastings Software Inc.). Design of olignucleotide primers was performed using Primer Designer Version 2.01 (Scientific & Educational Software). Prediciton of protein localization and predicition of cleavable signal peptides was performed using PSORT II (http://psort.nibb.ac.jp/form2.html) (Nakai and Kanehisa, 1992) or MitoProt (http://www.mips.biochem.mpg.de/cgi- bin/proj/medgen/mitofilter/) (Claros and Vincens, 1996). Predcition of coiled-coil

63 domains was performed at Swiss EMB net node server (http://www.ch.embnet.org/software/COILS_form.html) (Lupas et al., 1991). Generation of sequence contigs was performed using SeqMan Version 5.00 (DNASTAR Inc.) and protein alignments were performed using MegAlign Version 5.00 (DNASTAR Inc.). BLAST homolog searches were conducted using the BLAST server at the National Center for Biotechnology Information at the National Institutes of Health (http://www.ncbi.nlm.nih.gov/BLAST/) and Aspergillus and fungal searches genome searches were conducted using the BLAST servers available at the Broad Institute at the Massachusetts Institute of Technology (http://www.broad.mit.edu/annotation/).

64

Figure 2.1 Generation of a gene deletion construct by 3-way fusion PCR The Af-pyroA gene deletion cassette (A) is fused with fragments of the genome sequences upstream (B) and downstream (C) of the gene to be deleted (genX) by fusion PCR, forming a functional deletion construct (fusion PCR reaction with primers 5’fP3 and 3’rP6) . The fusion of the three individual fragments is dependant on the overlapping sequence incorporated in primers (n red). After transformation, proper replacement of genX by Af-pyroA is confirmed by diagnostic PCR using a primer set that targets sequences either side of this integrated construct (P7 and P8).

65

Af-pyroA A ΔP1 ΔP2 5’fP3 5’fP4 B >1.0Kb genX >1.0Kb C 3’fP5 3’fP6

A Af-pyroA 5’fP3 B C

3’fP6

Af-pyroA

Af-pyroA X X genX

P7 Af-pyroA P8

Figure 2.1

66

Figure 2.2 Generation of a C-terminal GFP tagging construct by 3-way fusion PCR The tagging cassette is made up of sequences encoding a flexible GA5 (5X glycine- alanine) linker followed by a sequence encoding a GFP polypeptide and an Af-pyrG auxotrophic marker (A). After individual PCR amplification of fragments A, B and C, due to the overlapping sequences incorporated into primers (D, E), A, B and C fuse into one construct via 3-way fusion PCR (F). Proper integration is confirmed by diagnostic PCR using a primer set that targets sequences either side of this integrated construct (G)

67

GA5-GFP Af-pyrG D A E

>1.0Kb D B genX C >1.0Kb E

F A F D E genX GA5-GFP Af-pyrG C B D E

genX GA5-GFP Af-pyrG

genX GA5-GFP Af-pyrG

genX

G G genX GA5-GFP Af-pyrG

Figure 2.2

68 Gene # Gene Region Primer Sequence AN2692 mcnA 5' fP3 LU69 TCGATACGGCGATATATGGAGACACGAACG 5 'rP4 LU70 CATCCCATAACCCCAACCCAGTACCGTCAT 3' fP5 LU71 TCCTGATACTGCTTCTGAGACTACCAGCGT 3' rP6 LU72 CGAAGGAATCTACGCAGGAGACCAAGAACG ΔP1 LU73 CGTTCGTGTCTCCATATATCGCCGTATCGATAACTCCGGTCAGGTCGATCATCC ΔP2 LU74 ACGCTGGTAGTCTCAGAAGCAGTATCAGGACTGATGCCAGCCTCTGAAGACAGC P7 LU75 TGTTGCCGAGTGGTTGGATG P8 LU76 AGGCATATGATCGCGCATTC AN8858 mcnB 5' fP3 LU78 ATGACCGATTGACGCGCTGCGGGTTGAGAC 5 'rP4 LU79 CCGCGGAGTTGACTTTTTATGGCTCGGAG 3' fP5 LU80 ACGCGGCCTTGACGCTTCATGCCTTGTTGC 3' rP6 LU81 ACATCAAGCCCTGAACCTCTATAACCTCCG ΔP1 LU82 GTCTCAACCCGCAGCGCGTCAATCGGTCATTAACTCCGGTCAGGTCGATCATCC ΔP2 LU83 GCAACAAGGCATGAAGCGTCAAGGCCGCGTCTGATGCCAGCCTCTGAAGACAGC P7 LU88 GCGTGGATACACTGATCT P8 LU89 CCAAGTCTCCGTGTCTAC AN2871 mcnC 5' fP3 LU27 TCTTGTGCATGCTGCTCTAGGAAGTTTGCC 5 'rP4 LU28 GGAGGAGAAATGTGATGGAGCTCGAAGGAG 3' fP5 LU29 ACGGGCCCAGATATGAAAACCGACTTTCTC 3' rP6 LU30 AACTACTACGGGAACACTGGACGAGGTGGC ΔP1 LU25 TCCTTCGAGCTCCATCACATTTCTCCTCCCTAACTCCGGTCAGGTCGATCATCC ΔP2 LU26 GCCACCTCGTCCAGTGTTCCCGTAGTAGTTCTGATGCCAGCCTCTGAAGACAGC P7 LU6 ATCCAGTACGGCCATAGA P8 LU9 TACGGAGTCAGCCACAAC Table 2.1. List of primers for gene deletion

69 Gene # Gene Region Primer Sequence AN2692 mcnA 5’ GFP/S-fP3 LU96 CGTGTCTCCATATATCGCCGTATCG 5’ GFP/S-rP4 LU97 AGTCGACACGCTGGTAGTCTCAGAAGCAGT 3’ GFP/S-fP5 LU98 TGAGCTATACAGCTATTCTGCTGCTGTAAG 3’ GFP/S-rP6 LU99 CTCAAGACGACACAACACAACCGAC GFP/S-P1 LU100 ACTGCTTCTGAGACTACCAGCGTGTCGACTGGAGCTGGTGCAGGCGCTGGAGCC GFP/S-P2 LU101 CTTACAGCAGCAGAATAGCTGTATAGCTCAGTCTGAGAGGAGGCACTGATGCG GFP/S-P7 LU75 TGTTGCCGAGTGGTTGGATG GFP/S-P8 LU103 GAATCTATGCCGGAGACC AN8858 mcnB 5’ GFP/S-fP3 LU90 CAGGATAGGACGACAACCATCAAGC 5’ GFP/S-rP4 LU91 AGGCCGCGTGTTGCTGCCGCGCTGAAAAGT 3’ GFP/S-fP5 LU92 TGACGCTTCATGCCTTGTTGCTTTATCTTC 3’ GFP/S-rP6 LU93 CAACCGACTGGATGTAACCTTCGAG GFP/S-P1 LU94 ACTTTTCAGCGCGGCAGCAACACGCGGCCTGGAGCTGGTGCAGGCGCTGGAGCC GFP/S-P2 LU95 GAAGATAAAGCAACAAGGCATGAAGCGTCAGTCTGAGAGGAGGCACTGATGCG GFP/S-P7 LU88 GCGTGGATACACTGATCT GFP/S-P8 LU102 GTCGACACAGTTCATTGC AN2871 mcnC 5’ GFP/S-fP3 LU40 GGAACACGCACGGAGAAGTCATC 5’ GFP/S-rP4 LU39 GTGACCATAGTTACCACCCCAGCCACCTCG 3’ GFP/S-fP5 LU38 TAAAGGATCAAGCTCTGATGTTCCGAGG 3’ GFP/S-rP6 LU41 CCGTGTATGGGAGCTTGGATAGACT GFP/S-P1 LU42 CGAGGTGGCTGGGGTGGTAACTATGGTCACGGAGCTGGTGCAGGCGCTGGAGCC GFP/S-P2 LU43 CCTCGGAACATCAGAGCTTGATCCTTTAGTCTGAGAGGAGGCACTGATGCG GFP/S-P7 LU51 GAGTCAGCCACAACTGCCGT GFP/S-P8 LU52 TTCTTGCGCGAGTGTTAAGC AN11145 CgrA 5’ mRFP/S-fP3 LU114 AAATCGAGTCTGGCCGTCAGGGTGAAGC 5’ mRFP/S-rP4 LU115 AGAGTTGAGGAGCTTGTTTCGTTTTTCTC 3’ mRFP/S-fP5 LU116 TGAACAATGTTATCAGTGTCAGTGTGC 3’ mRFP/S-rP6 LU117 GAATTCCTATCTTCCCTCCCCCCTCCTC mRFP/S-P1 LU118 GAGAAAAACGAAACAAGCTCCTCAACTCT GGAGCTGGTGCAGGCGCTGGAGCC mRFP/S-P2 LU119 GCACACTGACACTGATAACATTGTTC ACTGTCTGAGAGGAGGCACTGATGCG mRFP/S-P7 LU127 GCAGAAGACCGGACAATC mRFP/S-P8 LU128 CCGACATCCACGTGTATC Table 2.2. List of primers for gene tagging

70 Gene # Gene Region Primer Sequence AN0745 Fibrillarin 5’ GFP/S-fP3 LU129 TAGTAAGCTGGCTGCCGGTATTCTC 5’ GFP/S-rP4 LU130 CGACGAACGATTGTAGATACCGGAGACGAT 3’ GFP/S-fP5 LU131 TAAAAGCCAAAAGCAGGGTCCTTTATAACG 3’ GFP/S-rP6 LU132 GAAGATGACAGACGAATCGCAGAGG GFP/S-P1 LU133 ATCGTCTCCGGTATCTACAATCGTTCGTCG GGAGCTGGTGCAGGCGCTGGAGCC GFP/S-P2 LU134 CGTTATAAAGGACCCTGCTTTTGGCTTTTACTGTCTGAGAGGAGGCACTGATGCG GFP/S-P7 LU135 CATTGCGGTTGAGTCTCC GFP/S-P8 LU136 CGCCGGTAATAAGGACAG AN1367 Bop1 5’ GFP/S-fP3 LU138 ATCTTACCCAGTATCCCTTCCGTCG 5’ GFP/S-rP4 LU139 CATCCACAGCCGACACGTTCCATCCGCACC 3’ GFP/S-fP5 LU140 TAATTAGTCGATCTGCTGCATGCTTACTATC 3’ GFP/S-rP6 LU141 GCCGTCTGGAAACTCTTCTCTTCGT GFP/S-P1 LU142 GGTGCGGATGGAACGTGTCGGCTGTGGATG GGAGCTGGTGCAGGCGCTGGAGCC GFP/S-P2 LU143 GATAGTAAGCATGCAGCAGATCGACTAATTA CTGTCTGAGAGGAGGCACTGATGCG GFP/S-P7 LU144 AAGAGCACACCTCCTCAG GFP/S-P8 LU145 TTCTGAGGCATCAGCTCG AN3455 Nrap1 5’ GFP/S-fP3 LU146 CATCGACCCAGCAATGAATAGCGTG 5’ GFP/S-rP4 LU147 CTTTTCGTGGACTTCGATCCGAGAAACCAT 3’ GFP/S-fP5 LU148 TGAACGACATTATGTTGTTATATGTAGGTT 3’ GFP/S-rP6 LU149 TGGTGAGGCGAAGCTAGAGTTTGAG GFP/S-P1 LU150 ATGGTTTCTCGGATCGAAGTCCACGAAAAG GGAGCTGGTGCAGGCGCTGGAGCC GFP/S-P2 LU151 AACCTACATATAACAACATAATGTCGTTCA CTGTCTGAGAGGAGGCACTGATGCG GFP/S-P7 LU152 TTCCTCAGGACGTTGCAC GFP/S-P8 LU153 ACGATAACTGCCGTCAGC AN0410 BIMG 5’ GFP/S-fP3 LU154 TCCCACTGATGTAAGTCTCCAGGCG 5’ GFP/S-rP4 LU155 CTTCTTTTGCTTTCGCGGAGGAGTGATGGG 3’ GFP/S-fP5 LU156 TAAGCATACGAGCTTTTCCATGTTCAGCAG 3’ GFP/S-rP6 LU157 GCGCAGCGCACTCACAGACAAGAGG GFP/S-P1 LU158 CCCATCACTCCTCCGCGAAAGCAAAAGAAG GGAGCTGGTGCAGGCGCTGGAGCC GFP/S-P2 LU159 CTGCTGAACATGGAAAAGCTCGTATGCTTA CTGTCTGAGAGGAGGCACTGATGCG GFP/S-P7 LU160 CCATCGCGGCTATCATTG GFP/S-P8 LU161 GCGCGGTATAGAATCTTG AN0253 Topo I 5’ GFP/S-fP3 LU162 CAGATGATGCTCGACCTCGATCCGT 5’ GFP/S-rP4 LU163 AAACTCCCAGTTCTCATCGACGGACTTAAT 3’ GFP/S-fP5 LU164 TGATGAACTGCATTTGGCTTTTCCTGATTA 3’ GFP/S-rP6 LU165 GTCACTTGAAACGCGCTTAGACTGC GFP/S-P1 LU166 ATTAAGTCCGTCGATGAGAACTGGGAGTTT GGAGCTGGTGCAGGCGCTGGAGCC GFP/S-P2 LU167 TAATCAGGAAAAGCCAAATGCAGTTCATCA CTGTCTGAGAGGAGGCACTGATGCG GFP/S-P7 LU168 AAGATGAGCGAGCGAGTG GFP/S-P8 LU169 TTGGTCTGTTGAACGGAG Table 2.2. (continued) List of primers for gene tagging

71

CHAPTER 3

COPY NUMBER SUPPRESSORS OF THE NIMA1 MUTANT

3.1. Introduction The intrinsic power of genetic analysis is highlighted in its ability to allow one to define the precise biological function of a particular gene product and decipher the exact biochemical, physiological and developmental consequence to a cell in absence of that function. It also presents the possibility of isolating interesting mutants in the absence of any prior knowledge about the function that will be affected thereby leading to identification of new functions. While there are many different types of mutations, this section will deal solely with the isolation and identification of conditional mutants, mutants that produce their phenotypic effect only under certain conditions. In order to identify new genes that may interact with these mutations the concept of suppressors will be discussed as well. The effect of circumventing the phenotypic change of the mutation in the original gene is called suppression. There are different mechanisms for suppressing mutations in gene encoding for proteins. A secondary mutation in the same gene can sometimes compensate for a primary mutation giving intragenic suppression. Suppressor mutations in other genes, called extragenic suppressors, can be of two kinds: translational and physiological. Translational suppressors (amber, ochre, opal suppressor genes) compensate for the alteration within mRNA thus restoring the wild type phenotype while leaving the original mutation in DNA untouched. Physiological suppressors are usually mutations in genes functionally related to the original mutation, i.e. genes acting in the same pathway. A fourth type of suppression termed copy number suppression does not involve a second mutation but the addition of increasing number of copies of a particular

72 gene which compensates for the mutant function by physically changing the properties of the mutant protein or by changing the way it is regulated. The crux of this thesis is the application of copy number suppression to the mutant gene nimA1 as a genetic strategy to identify other nimA interacting proteins and decipher more about nimA regulation and the regulations of proteins that are targeted by nimA protein kinase.

3.1.1. Conditional mutants Formerly, the range of genetic analysis was limited by the fact that mutants defective in essential functions of a cell could not be analyzed as they could not be propagated. Conditional mutants, which display a characteristic mutant phenotype under some conditions (termed restrictive or non-permissive) but otherwise show wild type characteristics under other conditions (termed non-restrictive or permissive), provide a good solution to extend genetic analysis to many essential genes. The most widely studied conditional mutants are temperature sensitive, in which cells display a mutant phenotype at some temperatures but not at others (Horowitz and Leupold 1951). Other conditions may include change in pH or osmolarity. Extensive research on a variety of systems has amply demonstrated the utility of conditional temperature mutants. Some examples include, study of gene expression in bacteriophage lamda by identification of conditional lethal mutant of lamda genes (Herskowitz 1973), study of RNA synthesis in bacteria (Escherichia coli) by identification of conditional mutant aminoacyl ribonucleic acid synthetase (Eidlic and Neidhardt 1965), study of cell determination and fate in flies (Drosophila melanogaster) by identifying temperature sensitive mutants of the bithorax locus (Tasaka and Suzuki 1973) and the study of various cell cycle regulatory genes (cdc, nim, bim) by identifying cdc mutants in fungi (Hartwell, Culotti et al. 1974; Morris 1975; Nurse, Thuriaux et al. 1976). Temperature sensitive mutants (ts for short, their permissive and restrictive conditions are relatively low and high temperatures, respectively, within a normal growth range) are generally thought to be missense mutations in the structural genes for proteins (Carr and Kaguni 1996; Otterson, Modi et al. 1999; Smyth and Belote 1999). The resulting amino acid substitution partially destabilizes the protein products of the mutant

73 genes, resulting in their improper three dimensional structures and hence functions at higher temperatures (Maas and Davis 1952; Edgar and Lielausis 1964; Eidlic and Neidhardt 1965). In certain other cases, ts mutants may arise due to partial deletion or nonsense mutations near the ends of genes giving rising to truncated gene products or they may be ts for protein synthesis alone, meaning, that any gene product already present (made at permissive conditions) at the time of shift will support a limited period of continued growth. A conditional ts mutation bimG11 in one of the type 1 protein phosphatase genes bimG lies in the intron sequence of the gene that leads to an impaired splicing of the mutant mRNA at restrictive temperature. Its expression hence leads to the production of a truncated gene product that interferes with bimG function (Hughes, Arundhati et al. 1996). A variety of genetic analyses have made it clear that most genes of an organism have potential sites for ts mutations (Harris, Cheng et al. 1992). Because the defective gene products of ts mutants are frequently not fully functional at permissive temperatures or not fully non-functional at the restrictive temperatures, or both, the more useful ts mutants are ts lethals, with mutation in essential genes that cause discernible phenotypes.

3.1.2. Genetic screens and identification of conditional ts mutants A genetic screen is an experimental process for selecting individuals possessing a particular phenotype of interest. A screen designed to search for new genes is often referred to as forward genetics. The essence of any genetic screen or selection is to identify a process of interest, to predict the phenotype of a mutant unable to carry out that process and to devise a method to identify mutants with that phenotype. Genetic screens can be broadly classified into temperature sensitive screens and suppressor screens. Here we shall discuss the temperature sensitive screen using the identification of the nimA mutants as example and subsequently discuss the suppressor screen. A temperature sensitive screen is aimed at identifying conditional temperature dependent mutants. In these screens, cells are propagated at the permissive temperature and later screened at the restrictive temperature for the phenotype of interest. These are identified by replica plating, and looking for colonies that can grow under the permissive

74 temperature but not under the restrictive one. Once identified, the cells are then further examined for the desired phenotype. In many situations, it may be practical to obtain spontaneous ts mutations, however most temperature sensitive screens use additional mutagenic agents called “inducers” to increase the frequency of mutations. In early 1975, Ron Morris set up a mutagenic temperature sensitive screen using Aspergillus nidulans to identify cell cycle mutants (Morris 1975). Conidia (spores) were harvested and irradiated with ultraviolet light for random mutagenesis. The spores were then plated and allowed to grow at the permissive temperature. Four days later (approximate duration of time for spores to grow into colonies) surviving colonies were replica plated and incubated at both permissive and restrictive temperatures. Presumptive ts mutants, identified by their failure to grow at the restrictive temperature, were then grown and maintained at the permissive temperature. The percentage of survival was typically about 1% where as the yield of ts mutants were 3-5%. In order to classify the library of temperature sensitive mutants obtained, each mutant was scored for the desirable phenotypes, which were the presence of mitotic spindles and condensed chromatin to determine the spindle mitotic index (SMI) and the chromosomal mitotic index (CMI), respectively. Mutations resulting in the lowering of the mitotic indices were considered to affect normal entry into mitosis and termed nim mutants or “never-in-mitosis” mutants. Mutations causing an increase in the mitotic indices were considered defective in their passage through mitosis and termed bim mutants or “blocked-in-mitosis” mutants. Dominance tests (to determine if the mutation was recessive or dominant in the presence of a wild type allele) and complementation analysis (to determine if two independently isolated recessive mutations affected the same gene function), identified 9 bim mutants in 6 genes (2 alleles each of bimA, bimB and bimD) and 26 nim mutants in 23 genes (4 alleles of nimA). In addition to these mutants, mutants affecting septation (sep mutants) and nuclear migration within the cell (nud mutants) were also identified (Table 1.1). All these mutants showed defined cytological characteristics at the restrictive temperature representative due to mutations in various cell cycle specific genes. At permissive temperature however, they behaved just like wild type strains.

75 3.1.3. The nimA ts mutants At 32˚C, mitosis is a five minute process that occurs after a well defined G2 phase roughly lasting for 20 minutes. Nuclei in G2 are characterized by a lack of condensed DNA coupled with the presence of duplicated but non-separated spindle bodies located in the nuclear membrane. Progression into mitosis on the other hand is defined by condensation of DNA and the formation of an intranuclear spindle between two separated spindle poles (Oakley and Morris 1983; Osmani, Pu et al. 1988). Therefore, the transition of a nucleus from G2 to M is quiet dramatic and shows distinct morphologies for each phase. Strains carrying different mutant alleles of nimA (nimA1, nimA5 and nimA7) block in late G2 when grown at the restrictive temperature (Bergen, Upshall et al. 1984). If the blocked cells are returned to the permissive temperature, they enter mitosis in a synchronous manner. The duplicated spindle pole bodies separate, the DNA condense and spindles form in the next 5 minutes (Oakley and Morris 1983). DNA mediated complementation of the mutant phenotype of the nimA5 allele was done to clone the wild type gene. The molecular cloning of the G2-M specific gene, nimA by mutant rescue of nimA5 allowed further characterization of the gene (Osmani, May et al. 1987). To define the different mutant alleles of the nimA at the DNA level, DNA from a wild type strain along with the nimA1, nimA5 and nimA7 strain were amplified using polymerase chain reaction and sequenced. For each mutant allele, a point mutation causing a single amino acid substitution was identified (Refer Table 1). Each of these mutation maps to a different location of NIMA kinase (Figure 3.1) (Pu, Xu et al. 1995). The nimA7 mutation lies in the catalytic domain right next to a lysine residue shown to be essential for the kinase activity of NIMA. The nimA5 mutation maps to another distinct site in the catalytic domain as well. The nimA1 mutation on the other hand resides in the non-catalytic C-terminus and is in the same region as one of the two potential nuclear localization signals present in the C-terminal regulatory domain of NIMA. For all three mutants arrested in G2 at the restrictive temperature, the following results hold true: 1) The levels of NIMA protein are elevated in the cytoplasm (Pu, Xu et al. 1995; De Souza, Osmani et al. 2000; De Souza, Horn et al. 2003) and the protein expression is non affected 2) The accumulated protein is not hyperphosphorylated

76 therefore not fully activated but does get rapidly phosphorylated and located to the nucleus when released into permissive temperature 3) The kinase activity of the protein kinase CDK1-cyclin B is fully activated (Osmani, McGuire et al. 1991). The hyperphosphorylation of NIMA is seen after activation of CDK1-cyclinB during mitotic initiation (Ye, Xu et al. 1995). Since each of the nimA alleles causes a G2 arrest with fully activated CDK1 activity, it is unclear why NIMA is not hyperphosphorylated and therefore not fully functional at the arrest point. One possibility is that at the G2 arrest point, the NIMA protein is in a conformation unsuitable for CDK1-cyclin B driven hyperphosphorylation. A second possibility is that fully activated CDK1-cyclin B needs further phosphorylations to be able to phosphorylate NIMA and is unable to modify itself in the mutants. A third possibility is the lack of any interaction due to different sub- cellular localization of NIMA and CDK1-cyclin B prior to initiation of mitosis. While all the three different alleles of nimA at the restrictive temperature have identical biochemical features, it is important to distinguish between the three in terms of their functionality. The nimA5 and nimA7 mutations lie in the catalytic domain and therefore interfere with the kinase function of NIMA protein. Their basal level of kinase function is reduced to, and is equivalent to a kinase dead version of NIMA. The nimA1 mutation on the other hand does not disrupt the kinase domain and hence its kinase activity, though sub-optimal, is still partially intact. The nimA1 mutant however, still arrests in G2 due to the inability of the NIMA1 protein to localize to the nucleus where its function is required (De Souza, Horn et al. 2003). Very surprisingly, extragenic suppressors of nimA1 were identified to be mutations in nuclear pore complex proteins (suppressor of nimA1: sonA and sonB) that allowed for the nuclear entry of NIMA1 protein and its proper localization resulting in entry into mitosis (De Souza, Horn et al. 2003). These suppressor mutants however do not suppress the nimA5 or nimA7 phenotype, showing that basal activity of NIMA is required for the suppression. Moreover, the double mutants between the suppressor mutations in sonA and sonB and either nimA5 or nimA7, result in synthetic lethality (De Souza, Horn et al. 2003) most probably due to dominant negative effects of inactive nuclear NIMA5 and NIMA7 . In other words, a nimA5 or nimA7 mutant can only be suppressed if its kinase activity is

77 compensated. Suppression of a nimA1 mutant however should be possible through modification of its expression levels, its localization or through the modification of substrates that nimA regulates such that the requirement for full NIMA kinase activity can be lowered.

3.1.4. High copy number suppression of nimA1 – the AMA1 system 3.1.4.1. The utility of suppressor screens Aspergillus NIMA is essential for mitotic entry and its degradation is necessary for mitotic exit (Osmani, McGuire et al. 1991; Pu and Osmani 1995). The NIMA kinase regulates multiple events during mitotic progression. NIMA contributes to the timing of mitotic entry through controlling the localization and/or activation of the CDK1/cyclin B kinase (Wu, Osmani et al. 1998). Screening for extragenic suppressors of the nimA1 allele led to identification of two components of the nuclear pore complex, SONA, a homologue of yeast Gle2/Rae1, and SONB, a homologue of human Nup98 (De Souza, Horn et al. 2003). Genetic interactions suggest that NIMA might directly participate in the nuclear uptake of CDK1/cyclin B through the nuclear pore (Wu, Osmani et al. 1998; De Souza, Horn et al. 2003). NIMA is also strongly implicated in promoting chromatin condensation. Histone H3 is phosphorylated by NIMA on Ser-10 (De Souza, Osmani et al. 2000). Phosphorylation of H3 Ser-10 is closely correlated with chromatin condensation in many eukaryotes and this could explain why ectopic expression of NIMA drives premature chromatin condensation in yeast, Xenopus and human cells (O'Connell, Norbury et al. 1994; Lu and Hunter 1995). In addition, NIMA is required for the regulation of mitotic spindle formation (Osmani, Pu et al. 1988) and for the localization of CDK1/cyclin B to the spindle poles (Wu, Osmani et al. 1998). Unfortunately, the mechanisms by which NIMA kinases operates still remains poorly understood primarily because few direct substrates of NIMA have been identified. In addition, the NIMA profile during the cell cycle is very specific. NIMA protein and mRNA levels begin to increase in G2, reaching their peak at M (Osmani, May et al. 1987). At mitotic exit, there is a rapid decline in both protein and mRNA levels. The NIMA protein levels is rapidly degraded by the APC (Osmani and Ye 1996; Ye, Fincher

78 et al. 1998), however not much is known about the regulation of NIMA mRNA turnover. Also, the transcriptional activation at G2 and inactivation at mitotic exit remains unknown and thus presents an interesting topic of pursuit. A good way to get at regulatory aspects of NIMA function is to try identifying other proteins that interact with NIMA using genetic screens. The traditional extragenic suppressor screen for nimA1 utilized the concept of second site mutations to identify interacting genes of nimA but did not yield any other genes apart from the two specific interacting nucleoporins (De Souza, Horn et al. 2003). Hence, a second approach using a high copy number suppressor screen was designed to identify interacting genes of nimA. High copy number suppressors often show protein-protein interactions between the mutant and the suppressor gene product and therefore identify physically interacting proteins. As mentioned before, a high copy number suppressor is not a secondary mutation but instead the manipulation of gene copy number resulting in the over expression of genes. In order to perform a high copy number suppressor screen, two requirements have to be met. One, a set of DNA fragments which represents the entire genome of (a gene library) has to be available. And two, a system to introduce these clones in high numbers into the mutant strain to check for suppression. A gene library can be constructed by introducing sets of DNA fragments into circular cloning vehicles called plasmids. In a typical plasmid vector library the average size of the DNA fragments is 10- 12 Kb while the average gene size in Aspergillus is approximately 3 Kb. Therefore, each plasmid may carry about 3-4 genes. These genes can then be tested for suppression of the mutant phenotype by introducing the plasmids into the mutant strain and looking for reversion into wild type behavior at the restrictive condition. The gene copy can be manipulated by directed integration (gene targeting) of extra copies into the genome by recombination or by introducing them on vehicles that allow them to be maintained extrachromosomally, as episomes, without integrating into the genome. The process of introducing plasmids into a strain is called transformation. The frequency of correct gene targeting by homologous recombinations i.e. a chromosomal crossover between exact sequence matches is often low due to the high frequency of non- homologous integration of transforming DNA fragments.

79 For episomal transformation, this inefficiency of proper landing causes no problem as there is no need for genomic integration. The copy number of episomes per cell can be maintained easily by maintaining selection for the episomes. The advantage of introducing extra gene copies extrachromosomally on episomes rather than integrating them into the genome are as follows. The site of landing for the extra genomic copies does not need to be dealt with which avoids modifying the genome that may cause additive effect to the mutant phenotype. It also allows the task of re-isolation of the suppressor plasmid to be much easier.

3.1.4.2. The AMA1 plasmid With the exception of a few eukaryotes, the presence of chromosomal replicator (origin of replication) on a circular vector is not sufficient to maintain its replication episomaly. Successful eukaryotic episomal systems are based on naturally occurring eukaryotic plasmids such as the 2μm plasmid from S. cerevisiae (Beggs 1978) or on synthetic plasmids that utilize genomic elements located on chromosomes that are able to retain their ability to support replication extrachromosomally (Aleksenko and Clutterbuck 1997). An example of such sequence in S. cerevisiae is the autonomously replicating sequence (ARS) (Stinchcomb, Struhl et al. 1979). However, in organisms apart from the budding yeast the existence of ARS type DNA sequences have been difficult to find. In A. nidulans, the identification of an effective plasmid replicator sequence - AMA1 (Autonomously Maintained in Aspergillus) is therefore of special value. AMA1 was first identified due to the instability of transformant colonies with an gene bank caused due to sectorial plasmid loss (Gems, Johnstone et al. 1991). The chromosomal AMA1 sequence was shown to confer on plasmids the ability to persist in Aspergillus nuclei as circular super coiled molecules. The plasmids were structurally stable and did not undergo chromosomal integration (Gems, Johnstone et al. 1991). Synthetic AMA1 plasmids, bearing marker genes, indicated that gene expression was not generally affected by the episomal location and that the plasmid copy number only depended on the selective system used (Aleksenko and Clutterbuck 1996). The AMA1 system also had the advantage over integrative systems by having a much higher transformation rate. The

80 sequence resulted in an enhancement of transformation frequency of about 2000 fold compared to conventional integrating plasmids and the average number of plasmid copy number maintained per nucleus was 10-30 (Gems, Johnstone et al. 1991). The AMA1 system therefore provided a perfect experimental set up to perform a high copy number suppression screen. The AMA1 gene library is based upon the pRG3-AMA1-NotI vector, which includes the AMA1 sequence, the N. crassa nutritional marker pyr4, the antibiotic ampicillin resistant marker ampR and a multiple-cloning site with unique restriction enzyme sites. pyr4 allows for selection in A. nidulans while the ampR allows for selection in E. coli. The advantage of having two markers is that the AMA1 plasmid causing the suppression in can be re-isolated and propagated by subsequent transformation in E. coli. The same strategy was also used to make the gene library in the first place. The gene library was made by digestion of genomic DNA with six cutter restriction enzymes and subsequent cloning of these DNA fragments into the pRG3- AMA1 plasmid (Osherov and May 2000).

3.2. Results 3.2.1. Isolation of copy number suppressors – screen and experimental design A high copy number suppressor screen utilizing the AMA1 gene library was conducted for nimA1 (Figure 3.1). The nimA1 strain, LPW3, carries the nimA1 mutation and a mutation in the gene pyrG. pyrG encodes orotidine 5'-phosphate decarboxylase, the terminal enzyme in uridine 5'-phosphate biosynthesis (Oakley, Rinehart et al. 1987), required normally for regular growth and a mutation in pyrG (pyrG89) hence prevents growth in media lacking external sources of uridine. The AMA1 gene library vector, as mentioned earlier, carries an equivalent gene of the pyrG from the fungus Neurospora crassa called pyr4 (Osherov and May 2000). pyr4 is a functional homolog of pyrG but only shares minimal sequence similarity. Therefore, the successful transformation of the LPW3 strain by the AMA1 gene library plasmid can be selected by the complementation of the pyrG mutation by pyr4, on media that is lacking any external source of uridine/uracil.

81 LPW3 protoplasts (cells treated to remove their cell membrane so that they can receive external DNA) were transformed using different concentrations (1μg, 2μg and 4μg) of AMA1 library DNA and plated on uracil/uridine minus media and tested for growth at the restrictive temperature for nimA1, 42˚C. The plates were left in the incubator for at least 4 days after which the surviving colonies were retrieved. The surviving colonies were streaked three times to get clonal population of cells. Each of the colonies were then point inoculated on replica plates, along with the original nimA1 parent strain and a wild type strain but still carrying the auxotrophic pyrG marker, on media maintaining the nutritional selective pressure. The plates were then each incubated at 32˚C and 42˚C. Following observations were made regarding their growth: 1) The wild type grew at both temperatures while the nimA1 mutant only grew at 32˚C, 2) There were three degrees of growth for the potential suppressor colonies- those that grew as well as the wild type at both temperatures, those that grew intermediate to the wild type and nimA1 and those that grew at 32˚C but failed to survive at 42˚C. Note that some colonies failed to grow at both temperatures and were considered to have lost the plasmid due to instability. Colonies that were initially isolated from the screen but subsequently failed to grow at 42˚C were considered false positives.

3.2.1.1. Isolation of suppressor plasmids from suppressors In order to isolate the suppressing plasmids from the surviving colonies, total (genomic and plasmid) DNA was made from the suppressors. Spores from suppressor colonies grown on plates at 42˚C were used to inoculate liquid media in a single petri dish and further incubated at 42˚C for mycelial growth. After 24 hours, the mycelia was harvested and lyophilized. This material was then ground to a powder and used for a total DNA prep to get total DNA. Small volumes of the total DNA (10μL) was then used to transform DH5-α bacterial cells which were plated on LB plates with 50µg/ml concentration of the antibiotic ampicillin. Successfully transformed bacteria were selected by their ampicillin resistance and four bacterial colonies per DNA transformation were subsequently inoculated into liquid LB with ampicillin and grown

82 overnight at 37˚C to perform mini-preps to re-isolate the plasmid DNA. The group of four plasmid DNA isolates represented a single suppressor set. The next step was to analyze these re-isolated plasmids. The AMA1 plasmid carries many unique restriction sites, sequences that can be recognized and cleaved by restriction enzymes, in its multi-cloning site. The construction of the gene library is such that each cloned fragment (insert) is flanked by unique specific restriction sites, allowing the use of restriction enzymes KpnI and NotI to cleave the plasmid into the vector and the insert. Once the suppressor sets (four of each) was isolated, they were all individually digested with the indicted restriction enzymes and the digests run on a 0.8% agarose gel to record the insert size and visualize the pattern of digestion (Figure 3.2). If the insert itself contains no restriction sites for KpnI, NotI or SphI, the plasmid will generate two bands only, one for the vector and one for the insert. If the insert instead contains a restriction site for either of the enzymes within it, then it will show up as multiple bands on the gel. In figure 3.2 all digests gave the same size for the vector (~9.0kb) and each set of four plasmids showed identical patterns of digestion indicating they carried the same insert. The pattern of digestion varied from one suppressor set to another but at times they were also identical across different sets. The purpose of enzyme digestion of the plasmids therefore was to categorize each potential suppressor set into representative groups, based on their pattern of digestion. Note that sometimes in a single suppressor set, the DNA digest pattern varied, most probably due to the suppressed strains carrying more than one type of plasmid. A single representative plasmid of each group was then sent for sequencing with primers flanking the insert to get approximately 600-800 bases of sequence from either ends of the insert. These sequences were them compared at the NCBI sequence database to find other homologous sequences to see if we could zero in on any particular gene. In order to rule out a possibility that nimA1 is suppressed due to the presence of the wild type nimA gene present in the plasmids, a straight forward experiment as explained in the following section was designed.

83 3.1.2. Confirmation of non-nimA suppressors by transformation in the nimA5 strain The nimA5 strain contains a conditional ts mutation in the catalytic domain of the protein leading to complete inactivation of the kinase function at the restrictive temperature. The suppression of nimA5 strain is only possible by its transformation with the wild type nimA gene, which can completely compensate for the kinase function. Potential nimA1 suppressor plasmids that may contain the wild type nimA gene should therefore be able to suppress the nimA5 strain as well as nimA1. Therefore, every re- isolated plasmid was re-transformed into the nimA1 and the nimA5 strain at the same time. Replica sets of plate were incubated at 32˚C and 42˚C. Plasmids that resulted in the suppression and growth of both nimA1 and nimA5 strain at the restrictive temperature were identified to contain the wild type nimA gene and discarded (Figure 3.3). From the initial screen, 67 numbers of colonies surviving at 42˚C was isolated. 32 numbers were re-confirmed to be suppressors of nimA1 by point inoculation and 6 numbers of groups were made of identical digest patterns. Of these, 14 were identified as true suppressors and finally 3 plasmids: A13, C13 and D15 were found to be unique. Note that these numbers represent the combined results of four separate (A, B, C and D) but identical screens and that A13 was identified 12 independent times.

3.1.3. Sequence characterization and sub-cloning of nimA1 suppressors Initial sequence information for each of these plasmids was retrieved using primers flanking the insert giving approximately 600-800 bases of sequence from either ends. The sequence was then queried against the NCBI genome database to identify any known homologous gene sequence from other organisms. However, these short sequences gave no significant gene hits. Therefore, the full sequence of each plasmid insert was retrieved using one of the two methods. A13 was sequenced using the traditional primer walking method as when these experiments were completed the Aspergillus total genome sequencing project was yet to be completed. The sequence for the C13 and D15 were retrieved from the completed Aspergillus genome database at http://www.broad.mit.edu/annotation/genome/aspergillus_group/MultiHome.html by using the paired sequence obtained initially with the flanking primers as delimiters.

84 Each retrieved insert sequence was further translated in all six frames by using the GENERUNNER software and analyzed for putative open reading frames (ORFs). The insert sequence was then prepared for sub-cloning by dividing the DNA sequence into smaller DNA units which when translated contained at least one major ORF. Primers were designed flanking these subunits and the DNA sequence amplified by PCR. The primers were also designed to contain overhang sequences incorporating the restriction sites for the NotI and the KpnI enzyme, such that they could be used to clone back the sub-clones into the parent plasmid pRG3-AMA1. The PCR amplified products were gel purified, sequentially digested along with the pRG3 vector plasmid with the two restriction enzymes and then set up for ligation and subsequent transformation into E. coli. Sub-cloned plasmids were then purified from the bacterial cells using a mini prep protocol and used to transform back the nimA1 strain to look for suppression at the restrictive temperature. Sub-clones still able to suppress the nimA1 mutant phenotype were further investigated as described below.

3.1.4. The multi-copy number suppressor of nimA1 genes mcnA, mcnB and mcnC As shown in figure 3.4, 3.5 and 3.6 respectively, a single sub-clone from A13, C13 and D15 was able to suppress nimA1 as strongly as the full length inserts. They were hereafter renamed as multi copy number suppressors of nimA1 – mcnC, mcnB and mcnA. Initially for A13, two sub-clones were made, each containing a single ORF translating exactly to a 12.5kDa and a 70kDa putative protein. Both of these sub-clones were unable to rescue the nimA1 phenotype at 42˚C.The third sub-clone containing DNA sequence encompassing both ORFs was however able to suppress nimA1. There was a possibility that the complementing DNA sequence actually encoded for a single protein and that the presence of an intron resulted in the two putative ORFs. Therefore, 5’ and 3’ RACE was performed to obtain full length cDNA sequence for this gene. The cDNA sequence confirmed a single ORF with a single intron placed between the two putative ORFs. The gene is called mcnC which encodes a protein of 90kDa (Figure 3.4).

85 The sequence for D15, when queried against the NCBI genome database revealed a highly conserved transcription factor domain called the fork-head domain. The sequence for this domain fell into one large ORF encoding for a 78.7kDa putative protein. D15 was therefore sub-cloned such that it contained the sequence for this single ORF and was shown to suppress the nimA1 mutant phenotype. RACE analysis identified the cDNA sequence which encoded a single protein from a single ORF with no introns. The gene was termed mcnB (Figure 3.5). For C13, the insert was divided into three sub-clones, each containing single ORF encoding potential proteins of 81.9kDa, 64.5kDa and 27.5kDa. The sub-clone containing sequence for the putative 30kDa protein resulted in suppression of nimA1. RACE analysis revealed its cDNA to encode, in addition to the 30kDa ORF further coding sequences interrupted by two introns. Put together, the gene encodes for a protein of 56.6kDa and is termed mcnA (Figure 3.6).

3.1.5. The effect of over-expression of mcn genes on NIMA protein profile In wild type cells, NIMA protein and kinase activity has a distinct cell cycle profile (Osmani, May et al. 1987; Osmani, O'Donnell et al. 1991). In the G2 phase of the cell cycle, NIMA activity and protein levels are low. At the transition from G2 into mitosis, NIMA becomes phosphorylated and NIMA protein and kinase levels begin to rise. During mitosis, NIMA becomes further phosphorylated and as the cell cycle progresses into anaphase, NIMA is subjected to anaphase promoting complex (APC) dependent ubiquitination and subsequent proteolysis in a manner that is dependent on the C-terminal PEST sequences. The APC however remains inactivated if the spindle assembly checkpoint (SAC) is engaged. Therefore, if a cell is arrested in mitosis by the artificial engagement of the spindle assembly checkpoint, the APC driven protein degradation of NIMA can be inhibited and the mitotic levels of NIMA measured (Figure 3.7). Such an arrest can be brought about by the use of certain drugs like Nocodazole and Benomyl. Both Nocodazole and Benomyl are microtubule depolymerising drugs and cause the spindle assembly checkpoint to get engaged due to the presence of unattached

86 kinetochores when cells enter mitosis (Osmani, May et al. 1987; Yu 2002). Such an arrest in mitosis due to artificial induction of the checkpoint is called a pseudo mitotic arrest. Over expression of the mcn genes using the high copy number plasmid AMA1 suppresses the conditional nimA1 mutant. This suppression, as mentioned before, may be due to modification of NIMA protein expression. To study if the over expression of mcn genes results in the up-regulation of NIMA1 protein, the following experiment was set up. The strains nimA1+empty vector-AMA1 (negative control), nimA1+WT nimA- AMA1 (positive control) and nimA1+mcn genes-AMA1 were grown to harvest spores. The spores were then inoculated at 2x106spores/ml concentration into three independent flasks containing rich media with no uridine/uracil to maintain selection for the AMA1 plasmid. The spores were allowed to germinate at the permissive temperature 32˚C till they reached the 8 to 16 nuclei stage at which point they were shifted to the restrictive temperature 42˚C by rapid shift method and incubated in the presence of the drug Nocodazole. After 3 hours at 42˚C the cells were harvested. Protein preps were made from all three harvested strains and western blot analysis performed with NIMA specific antibodies to check for NIMA protein levels (Figure 3.8). The arrest point for each strain at the restrictive temperature may be one of the following. A G2 arrest if there is no suppression of nimA1 and cells exhibit the nimA1 mutant phenotype or an arrest in M, if there is suppression of nimA1 resulting in cells entering mitosis but unable to pass through it due to the presence of Nocodazole. Figure 3.8 is a western blot showing the levels of NIMA protein for each strain at its arrest point. Lane 1 is negative control, incapable of any suppression and hence representative of basal levels of NIMA. Lane 2 is positive control having over expressed wild type NIMA resulting in suppression of nimA1 hence showing a much stronger signal for NIMA. Lane 3, 4 and 5 shows the expression level of NIMA from the strain suppressed for nimA1 due to over expression of mcnC, mcnB, mcnA. The NIMA profile shows that over expression of mcnC does not cause the up-regulation of NIMA while over expression of mcnB and mcnA results in significant up-regulation of NIMA protein.

87 3.3. Discussion The identification of three genes that specifically suppress the nimA1 mutant but not the nimA5 mutant at the restrictive temperature, suggests that the suppressors are not genes with functions that directly replace the kinase function of NIMA protein or bypass suppressors. Given that the difference between NIMA1 and NIMA5 protein is the level of their activity, NIMA1 is capable of partial kinase function while NIMA5 is kinase dead (Pu, Xu et al. 1995; Ye, Xu et al. 1995; Wu, Osmani et al. 1998), suppressors that compensate for nimA1 function by being over expressed must result in some sort of modification to the properties of nimA1 gene expression and/or its gene product. In this regard, there are two possible mechanisms for the high copy number suppression of nimA1, as mentioned below.

3.3.1. nimA1 suppression by over expression of NIMA1 protein levels nimA1 cells at the restrictive condition are arrested in G2 with cytoplasmic accumulation of NIMA1 protein (De Souza, Horn et al. 2003). The cells are thus unable to enter mitosis due to the inability of NIMA1 protein to accumulate in the nucleus where its functions are required. It is assumed that the nimA1 mutation may lead to a conformation defect affecting the function of its nuclear localization signal; thereby preventing it from being translocated to the nucleus (Wu, Osmani et al. 1998). Even if a certain fraction of NIMA1 can enter the nucleus, it will not be expected to accumulate in sufficient amounts to carry out the mitotic function of NIMA1. On the other hand, if a copy number suppressor results in the overall up-regulation of NIMA1 protein levels and leads to a higher amount of NIMA1 protein being accumulated in the cytoplasm and by a modified transport mechanism, this may then result in a larger amount of NIMA1 entering the nucleus that matches the required nuclear levels resulting in entry into mitosis. Higher NIMA1 levels may also compensate for the partial kinase function leading to modification of other proteins that are responsible for localizing NIMA1 into the nucleus. The effect on protein expression may be transcription dependent or transcription independent and may be through increased translation or decreased degradation rate of NIMA1 protein or mRNA. Note that this potential mode of nimA1

88 suppression would not be expected to bring back full NIMA function and that the mcn suppressors were able to only partially suppress the temperature sensitivity of nimA1.

3.3.2. nimA1 suppression by modification of NIMA1 protein localization The copy number suppressors of nimA1 may not result in the up regulation of NIMA1 protein. In such cases, a second mechanism of suppression must function to translocate the cytoplasmic NIMA1 protein to the nucleus without modifying nimA1 protein expression. This is possible if over expression of a copy number suppressor gene results in the modification of the transport system such that the basal levels of NIMA1 can now enter the nucleus. The identification of two nuclear pore proteins, sonA1 and sonB1, as extragenic suppressor of nimA1 and not nimA5 adds value to this possible mechanism. Both sonA1 and sonB1 mutation relieve the G2 nimA1 arrest in part by allowing NIMA1 into the nucleus (De Souza, Horn et al. 2003). Therefore, a similar scenario can suppress the nimA1 mutant phenotype if the over expressed suppressor gene modifies the transport property of NIMA1 by affecting nuclear pore proteins like sonA/sonB or others. The alternative possibility is the modification of the import capacity of the nucleus by over expressing certain importins, proteins that may assist in carrying NIMA into the nucleus. The results from this section clearly shows that the over expression of mcnA and mcnB results in the up-regulation of NIMA1 protein, suggesting that a change in NIMA protein expression is responsible for the suppression of nimA1. On the other hand, mcnC over expression appears not to affect NIMA1 protein expression and thus must have an alternate mechanism for translocation of NIMA1 into the nucleus and cause the suppression of nimA1. Specific studies of each of these copy number suppressors will be discussed in the chapters to follow.

89 MCS (EcoRI, SacI, HIII, KpnI, N. crassa pyr4 BamHI, SmaI, NotI, SphI) (2.3Kb)

pRG3-AMA1- NotI AMA1 AmpR (2.1 Kb)

Figure 3.1. Plasmid map of vector pRG3-AMA1-Not1 The plasmid was used to generate the A. nidulans gene library. The plasmid contains the A. nidulans AMA1 sequence, the ampicillin resistance gene, AmpR from E. coli and the N. crassa pyr4 marker. The multi-cloning site (MCS) contains the unique RE sites KpnI, BamHI, SphI and NotI. A. nidulans total genome digested with RE SauIIIA was cloned into the BamHI site present in the multi-cloning site (MCS). The vector is approximately 9.0kb in size and contains average inserts of 6-6.5kb of A. nidulans gene fragments. KpnI and NotI were used to retrieve the inserts.

90 1. Transformation of nimA1 strain with pRG3-AMA1-NotI containing A. nidulans gene library

2. Replica spot transformants at 32˚C and 42˚C, select those that partially complement the nimA1 phenotype at 42˚

3. Re-confirm partial suppression of potential suppressors at 42˚C

nimA1 nimA1+ nimA1+copy WT nimA number suppressors

Figure 3.2. Outline of copy number suppressor screen of nimA1 Step 1 involves transformation of strain LPW3 (nimA1) with pRG3-AMA1-Not I A. nidulans gene library and selecting transformants at 42˚C on MAG plates. Step 2 is replica spot of all transformants at 32˚C and 42˚C (shown in picture with controls), arrowheads point to potential nimA1 suppressed colonies. Step 3 is re-confirmation of potential suppressors by point inoculation at 42˚C (shown in picture with controls).

91

Figure 3.3. Isolation, purification & grouping of copy number suppressor plasmids. Picture shows restriction digestion with enzymes KpnI and NotI of copy number suppressor plasmids after isolation from E. coli. Each number represents a single suppressor set. Each lane shows the vector (9.0kb) and inserts bands. Multiple bands mean presence of KpnI or NotI site within the insert in addition to those in the plasmid vector. The digest patterns for groups 2, 4 and 5 are identical and were therefore grouped into a single complementation group.

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Figure 3.4. Confirmation of non-nimA suppressors by transformation in nimA5 Copy number suppressor candidates were re-transformed in both nimA5 and nimA1 conditional mutant strains and tested for growth at 32˚C and 42˚C. Transformed colonies of nimA5 and nimA1 both grew at the permissive temperature 32˚C (panels A and C). Suppressors other than WT nimA alone could complement nimA1 at 42˚C (panel D) but failed to suppress nimA5 at 42˚C (panel B). Suppressors unable to complement nimA5 at 42˚C (panel B) were confirmed to contain suppressor gene other than wild type nimA.

93

Figure 3.5. Sub-cloning of A13 Full length and sub-clones of A13 in pRG3-AMA1-NotI vector were re-transformed in nimA1 and tested for suppression at 42˚C. Sub-clone 3 suppresses nimA1 and RACE confirms sub-clone 3 to encode for a 90kDa protein. The gene was termed mcnC for multi-copy number suppressor of nimA1.

94

Figure 3.6. Sub-cloning of D15 Full length and a single sub-clone of D15 in pRG3-AMA1-NotI vector was re- transformed in nimA1 and tested for suppression at 42˚C. Sub-clone 1, as shown suppresses nimA1 and RACE confirms sub-clone 1 to encode for a 78.7kDa protein. The gene is termed mcnB for multi-copy number of nimA1.

95

Figure 3.7. Sub-cloning of C13 Full length and sub-clones of C13 in pRG3-AMA1-NotI vector were re-transformed in nimA1 and tested for suppression at 42˚C. Sub-clone 3 suppresses nimA1 and RACE confirms sub-clone 3 to encode for a 56.6kDa protein. The gene is termed mcnA for multi-copy number suppressor of nimA1.

96 β-tubulin

Figure 3.8. Western blot analysis for NIMA1 protein in copy number suppressed nimA1 strains nimA1 strain (LPW3) transformed with AMA1-plasmid containing wild type nimA, no insert, mcnC, mcnB and mcnA were processed for levels of NIMA1 protein at 42˚C during a mitotic arrest caused by the drug Nocodazole. Levels of tubulin are used as loading control. The non-specific band (arrow) may also be used as loading control. Lane with + empty vector represents basal levels of endogenous NIMA1 protein. Expression of wild type nimA, mcnB and mcnA from AMA1 plasmids results in up-regulation of NIMA1 levels. Similar expression of mcnC does not.

97

CHAPTER 4

MCNC GENETICALLY INTERACTS WITH NIMA

4.1. Introduction 4.1.1. NIMA interacting proteins Over expression of mcnC relieves the conditional G2 nimA1 arrest but fails to suppress the nimA5 mutant phenotype. In addition, mcnC over expression does not modify the NIMA1 expression profile (Figure 3.8). The data therefore suggests that mcnC may suppress the mutant phenotype of nimA1 by modifying the sub-cellular localization of NIMA1 and possibly other mitotic regulators rather than affecting NIMA protein expression. Screening for extragenic suppressors of the nimA1 allele has already identified two components of the nuclear pore complex, SONA, a homologue of yeast Gle2/Rae1, and SONB, a homologue of human Nup98 (De Souza, Horn et al. 2003).Genetic interactions and localization studies using the sonA and sonB mutants show the localization of NIMA1 and CDK1/cyclin B to be modified through the nuclear pore (De Souza, Horn et al. 2003). The suppression of nimA1 by modulation of NIMA1 localization through interaction between NIMA kinase and proteins that are directly involved with transport is an elegant way to regulate NIMA. Other nimA interacting proteins like TINA and TINC, which were identified as two hybrid interactors of nimA through a two-hybrid screen, have added valuable credence to NIMA function at additional sub-cellular locales. NIMA is known to play a role in spindle formation (Osmani, Engle et al. 1988) during mitosis and TINA was shown to regulate microtubule nucleation at the spindle poles (Osmani, Davies et al. 2003).

98 Hence, the advantages of identifying genes that interact with nimA are two fold. One, it helps to find other players in NIMA controlled pathways and add more understanding to the regulation of those particular pathway. Two, by characterizing nimA interacting genes, we also determine additional areas where NIMA protein kinase function is required. All the previous data about NIMA and NIMA interacting proteins have clearly established the role of NIMA during mitosis. NIMA accumulation is not only required for mitotic entry but degradation of NIMA at mitotic entry is also essential. Both these steps are very tightly regulated with additional roles played by the NIMA interacting proteins. More recently, preliminary studies of NIMA-GFP localization, has located the protein at septa (the cross walls present in growing germlings) and at the germ tip (the growing end of the germling), suggestive of NIMA function in controlling key aspects of A. nidulans cell morphogenesis.

4.2. Results 4.2.1. mcnC sequence characterization As a first step towards characterizing mcnC, the protein sequence was analyzed. mcnC encodes a 90kDa protein. There are no discernible domains found in the protein except a C-terminus domain with sparsely distributed FG repeats (7 of them) (Figure 4.1). FG repeats are typically the hallmarks of the FG family of nucleoporins that contain extensive regions of FG repeats, typically 200 -700 amino acids in length, which facilitate the passage of karyopherin-cargo complexes through the nuclear pore complex (Denning, Patel et al. 2003; Strawn, Shen et al. 2004). The typical number of FG repeats in FG nucleoporins range from 8 FGs (NUP49) to 53 FGs (SONB) (Osmani, Davies et al. 2006), both in their N-terminus and both proteins localize to the nuclear periphery demonstrating they are bonafide nucleoporins. The low number of FG repeats and its presence in the C-terminus of MCNC may mean that they are not of significant function to MCNC. Using mcnC protein sequence the BLAST assembled genome at the NCBI database was queried using blastp (Altschul, Madden et al. 1997) to identify putative homologous proteins from other organisms. Proteins sharing significant homology were identified in all eight Aspergilli species along with additional proteins in Neurospora

99 crassa, Schizosaccharomyces pombe and Ustilago maydis (Table 4.1). Sequence alignment done using CLUSTALW at http://www.ebi.ac.uk/Tools/clustalw/index.html shows the positions of the 7FG repeats to be conserved in the homologs of the Aspergilli.

4.2.2. MCNC protein is non-uniformly localized to the cytoplasm To determine whether MCNC localized to the nuclear periphery and constituted a nucleoporin, mcnC was endogenously tagged with green fluorescent protein at its C- terminus using the fusion PCR technique. Living cells expressing MCNC-GFP were visualized using confocal microscopy (Figure 4.2). MCNC-GFP was observed throughout the cytoplasm. DAPI staining of fixed cells revealed that areas lacking MCNC-GFP corresponded to nuclei. The cytoplasmic distribution of MCNC-GFP was not uniform but more structured. When cells were visualized through the cell cycle, the cytoplasmic signal with no nuclear signal was seen to be specific for interphase cells. As cells underwent mitosis, MCNC-GFP entered the nucleus and was visualized throughout the cell. A C-terminally tagged MCNC-GFP was also exogenously expressed under the control of the alcA promoter and an identical pattern of MCNC localization observed.

4.2.3. MCNC protein runs during SDS PAGE at a higher size than predicted While confirming expression of endogenously tagged MCNC-GFP by western blotting, it was noticed that the protein was running at a size much higher size (200KDa), almost double, than predicted (110kDa) (Figure 4.3A). This presented us with the possibility that the predicted protein size may be wrong due to the misidentification of the starting ATG codon due to automated gene calling and in reality the ORF may instead be a larger ORF giving the corresponding larger protein size. To test this possibility, protein made from the exogenously tagged AlcA-MCNC-GFP strain after induction for 0, 1 and 2 hours using threonine was analyzed (Figure 4.3B) and compared to the endogenously tagged MCNC-GFP by western blot using a GFP antibody. The logic for this was that since the AlcA version was constructed using a predefined ORF it should give us the expected 110kDa protein size whereas the endogenous tagging of mcnC was at the 3’ end and therefore if the actual ORF began at an upstream ATG, the endogenously tagged

100 version should reflect this by making a protein larger than 110kDa. In both cases, the size of MCNC protein was 200kDa suggesting that the protein was running at a higher size due to some modification to protein property rather than miscalculation of its predicted protein size. 3’RACE was also performed which confirmed the C-terminal end of the mcnC cDNA.

4.2.4. mcnC function is required for normal growth A null allele of mcnC gene was generated using homologous integration to replace its coding region with the A. fumigatus pyroA nutritional marker. The deletion construct was generated using fusion PCR (Yang, Ukil et al. 2004) and transformed into the wild type GR5 strain. Homologous integrants were confirmed by diagnostic PCR and Southern blot analysis. Deleted transformants were checked for heterokaryon formation to assess for mcnC essentiality. mcnC deletion (ΔmcnC) did not lead to heterokaryon formation and hence mcnC is not an essential gene. Further characterization of ΔmcnC was performed by testing it under an array of growth conditions as mentioned in section 2.11 to uncover potential conditional phenotypes. Deletion of mcnC caused restricted growth on solid media at both 32˚C and 42˚C when compared to appropriate control strains (Figure 4.4). The restricted growth was more pronounced at the elevated temperature indicating that ΔmcnC causes some temperature sensitivity. In addition to the growth tests, ΔmcnC was also self crossed to determine if the deletion leads to self- sterility. ΔmcnC was capable of self crossing and hence does not cause sterility.

4.2.5. Characterization of the ΔmcnC growth defects To characterize the type of defect causing the restricted growth of mcnCΔ, spores from the deleted strain were grown on selective media, fixed and stained by DAPI. This allowed visualization of general nuclear structure (Figure 4.5B). Bright field images of the same cells allowed visualization of general growth features (Figure 4.5B). During germination, spores break dormancy and undergo an initial isotropic growth followed by highly polarized growth, linked to rounds of para-synchronous nuclear divisions. The time of emergence of a germtube is usually after the first round of mitotic division when

101 two nuclei are present. Typical DAPI-stained nuclei are round to ovoid in shape and show the nucleolus as a less brightly stained shadow area. On germination on selective media at 42˚C for 7 hours, a control wild type strain showed 16 normally structured nuclei within a polarized germtube (Figure 4.5A). ΔmcnC spores on the other hand did not undergo germtube emergence similar to wild type strain spores, though the number of nuclei were the same. This indicates a defect in polarized growth which helps explain the reduced growth of ΔmcnC strains. Repeat experiments were done by growing both the control and ΔmcnC for different lengths of time. Note that ΔmcnC does not result in the prevention of polarization altogether, but instead causes a delay, as cells from later time point have germtubes. The length of these germtubes are however smaller in length than the control.

4.2.6. mcnC over expression affects polarity of the cells In order to be able to conditionally manipulate mcnC expression, mcnC was cloned into a plasmid containing the inducible alcA promoter (alcohol dehydrogenase I). The plasmid used, pAL5, is based on pUC19 and includes the N. crassa nutritional marker pyr4 and a multi cloning site downstream of the alcA promoter, followed by a 3’ 3.8kb fragment downstream of the H2A gene. The flanking H2A sequence allows for site-specific recombination and 3’ processing of cDNA inserts. Genes cloned into this site were demonstrated to be under the control of the alcA promoter at the level of transcription when introduced in (Waring, May et al. 1989). alcA based promoter expression is transcriptionally repressed in the presence of glucose (Bailey and Arst 1975; Lockington, Sealy-Lewis et al. 1985) and induced in the presence of ethanol (include ref in endnote-Creaser et al. 1985). alcA expression can also be manipulated in liquid media containing minimal yeast extract lactose (non-repressing non-inducing) by the addition of varying concentrations of threonine (inducing). Wild type GR5 strain was transformed with the alcA-mcnC plasmid. On transformation, the plasmid can integrate either at the endogenous mcnC locus, the alcA locus or the H2A locus and can do so in single or multiple copies. Successful transformants were replica spotted on both repressive (minimal media with glucose) and

102 inductive (minimal media with ethanol) media. Colonies grew to be healthy on repressive media but looked sickly on inductive media (Figure 4.6). Therefore, over expression of mcnC has deleterious effect on wild type cells. In order to characterize this effect, alcA- mcnC spores and alcA-mcnC-GFP spores were germinated in minimal media containing ethanol at 32˚C. After 10 hours, cells from both strains were visualized. Bright field images and GFP images were then documented (Figure 4.7). As shown in figure 4.7, the mcnC over expressed cells had atypical looking germlings with multiple germ tubes of varying lengths representative of multiples sites of polarization.

4.2.7. MCNC over expression affects nuclear pore protein localization A strain with green fluorescently tagged sonB (CDS641) was transformed with the alcA-mcnC plasmid. Spore from alcA-mcnC + sonB-GFP were germinated in minimal media containing ethanol at 32˚C. This allowed the visualization of the effect of over expressed mcnC on the nucleoporin SONB. After 10 hours, cells were visualized by confocal microscopy. Bright field images and fluorescent images were then documented (Figure 4.8). As shown in figure 4.8, the mcnC over expressed cells had the same multiple germ tube phenotype as described above along with dispersed SONB-GFP throughout the cytoplasm in some cells. In figure 4.8, one can correlate the presence of multiple germtubes with dispersion of SONB. Cells that did not show the dispersed SONB phenotype had SONB around the nuclear periphery as normal in wild type cells. This effect may mean that the expression levels of the AlcA construct were higher in cells having the multiple germtubes and hence were representative of highly over expressed mcnC cells.

4.2.8 mcnC genetically interacts with nimA In order to check if the ΔmcnC allele genetically interacts with nimA, we made double mutants of nimA (nimA1 and nimA7) with ΔmcnC and tested for enhanced temperature sensitivity. We predicted that the ΔmcnC deletion should make the nimA1 temperature sensitivity phenotype at 42˚C worse if it plays a positive role in NIMA regulation as predicted by the copy number suppression of nimA by mcnC. As can be

103 seen from Figure 4.9A, the mcnCΔnimA1 double mutants are temperature sensitive at 37˚C, a temperature at which both the single mutants are able to grow to wild type levels. We repeated the experiment with a different mutant version of nimA, nimA7, a mutation in the kinase domain of nimA resulting in a kinase dead version of protein at 42˚C. We found that ΔmcnC also showed synthetic genetic interaction with nimA7 at 34˚C (Figure 4.9B).

4.3 Discussion 4.3.1. mcnC over expression affects the nuclear pore complex protein SONB In mcnC suppressed nimA1 cells, the over expression of mcnC using AMA1 plasmid does not result in the up-regulation of NIMA1 protein. Instead, these strains show an effect of mcnC over expression on the nucleopore protein SONB. In wild type cells, SONB-GFP shows a characteristic nuclear pore complex localization at the nuclear periphery surrounding the nucleus during interphase. However SONB is known to disperse throughout the cell during mitosis. This is thought to be an essential step for mitosis to proceed. In cells with visible MCNC over expression (these cells had multiple germtubes) SONB-GFP was dispersed through out the cell for extended periods of time and did not show distinctive nuclear periphery localization. This data indicates that mcnC may directly participate in the nuclear uptake of cytoplasmic NIMA1 by promoting the release of SONB from the NPC, which is an essential step to overcome the nimA1 G2 arrest phenotype. This hypothesis is similar in principle to the mode of extragenic suppression of nimA1 by sonA1 and sonB1. The mechanism is thought to involve the phosphorylation of SONB by NIMA resulting in its dispersal from the nuclear pore (De Souza, Horn et al. 2003) and thereby allowing nuclear accumulation of essential mitotic regulators. mcnC may be a part of the same pathway or play a role in nuclear pore complex regulation at mitosis in an independent pathway.

104 4.3.2. mcnC genetically interacts with nimA The deletion of mcnC causes temperature sensitivity at 42˚C and leads to a delay in polarized growth. Interestingly, mutant nimA1 display synthetic lethality with ΔmcnC. Cells with nimA1 mutation show more severe temperature sensitivity when combined with ΔmcnC. This interaction in not allele specific as the double mutant nimA7 with ΔmcnC also causes synthetically lethality. The non-allelic specific interaction of mcnC with nimA mutants strongly indicates that mcnC may be a positive regulator of nimA. Hence mcnC inactivation leads to the ts nimA mutants becoming sicker. It cannot mean a functional overlap of the two genes as mcnC is unable to copy number suppress the nimA7 mutation. In addition, because extra mcnC does not cause elevated levels of NIMA to accumulate, the role of MCNC in the positive regulation of NIMA occurs post translationally. Collectively the data support the hypothesis that MCNA plays a positive role in the nuclear localization of NIMA during entry into mitosis.

4.3.3. Positive regulation of cell growth by mcnC When inoculated in appropriate growth media Aspergillus nidulans spores undergo an initial phase of isotropic expansion followed by polarized growth at a single point forming germlings with a single germtube. The initiation of germtube formation is closely related to the nuclear division cycle in A. nidulans. Cells that have undergone two rounds of nuclear division almost always show polarization. In mcnC deleted cells, 43% of cells at 4 nuclei stage showed no signs of a germtube as compared to 0% of wild type control. However, this effect was not permanent as these cells recovered at a later time point and became polarized. Therefore, ΔmcnC causes a delay in polarized growth and not a permanent defect. More surprisingly, when mcnC is over expressed it leads to an over production of germtubes. That the delay in polarization due to lack of mcnC and the increase in sites of polarization due to over expression of mcnC may be due to the positive regulation of mcnC by nimA is further supported by the recent localization study of NIMA protein. NIMA is seen to be localized to the septa and the germ tip. mcnC therefore may play a positive role in promoting polarized growth of A. nidulans through the regulation of nimA at these new sub-cellular localizations. The identification of mcnC

105 therefore has identified a NIMA interacting protein that may help us implicate NIMA function in a hitherto unknown area and may help unravel regulation pertaining to polar growth and how this may be regulated via the cell cycle.

106 mcnC (length 870) Organism Length Identity % E value Aspergillus fumigatus 905 71 0.0 Aspergillus oryzae 902 71 0.0 Aspergillus clavatus 908 68 0.0 Neosartorya fischeri 901 72 0.0 Aspergillus flavus 896 72 0.0 Aspergillus niger 780 67 0.0 Aspergillus terreus 794 58 0.0 Coccidioides immitis 871 56 2e-149 Neurospora crassa 1062 39 1e-136 Saccharomyces pombe 963 23 3e-12 Ustilago maydis 964 27 6e-08

Table 4.1. mcnC homologs mcnC encodes for a 870 amino acid containing protein. Blastp homology searches using mcnC identified proteins of similar sizes sharing significant homology in all members of the Aspergilli family and in the fungus Coccidioides immitis. Additional homologs were also found in Neurospora crassa, Schizosaccharomyces pombe and Ustilago maydis.

107 MSEVQSRSSASRGRVSARGGRGGYSSRGGRGGSRSTKPDVTEPTYEDEGELGQ MKKKYSDTLPMLKELFPDWTDEDLVFALEDADGDLEQAIDRISEGNVSQWGEV KKKTTDRSRPKPKEAQSTPTESATTAVRPGRGRGGFEGRGRARGDRGRGGRGG RAGTHANGTRTEKSSLPAEITPIADSATTTTATSSAETAAPETVSTTKDTPAM PEGTKKGWASLFAKPAVPPPQKKPAAPAPAPATAPPPAAPATEEPAPEPQPEE KPAEPDAAPAPAPAPAPVPVAVPLPADRAPQPAVPQPREEPQKATPTSADVSP AKDDLTKNNLAQIPDVSPPVPSATAASTVGSTVEPSAAATSTPARPTASALPT SAFKQNIRTPGTQRRVMEQQEAVVMPGNHAVDRAAVQFGSMGLNGEAADVDID ENREDTETRAQPPQHSPVAPRASLPPSTQAQAPPEAAAVSRPAPGLPPVPQAT AAENTFSDFARYDSQKPYDPFTQPLTQPQPQVQEPFANQAPVQPTVTTGSEYS PFYAGDQRLPYNYYNAYGQSQDASLAQRAAGFGVSGAEAQPQIPTTQPPTRYG HVEAPNSGHTTPNPTLPGVTQTPAAHHMPTQGAHAYGYGYPYYSNPHYASYMS QQYGRNRPIYDDARRYEDQYMPHSSQYGYGSQYGPYGKGGMYGQPHGFSYDHS ASPATAGSFNQGIPGRDSVYGRTGSAQPSESQQSAAGASAFGTGMTDVFGRSQ AGFGQNQPIAQQTPVSSEETKAFDASKTGGPSPSLSQANRPGSATNTPGQSQS QTGLPPLQGQQTQQGFGGYPHLNPQYGGLGGLGGHQTAANQTHHQATGYGNYG GAGFGNYYGNTGRGGWGGNYGH#

Figure 4.1. MCNC protein sequence mcnC encodes for a 870 amino acid protein with a molecular weight of 90kDa. The protein does not contain any distinctive domains but contains 7 dispersed FG repeats in the C-terminus, shown in red. FG repeats are hallmarks of nuclear pore complex proteins. The typical number of FG repeats in A. nidulans FG nucleopore proteins range from 8 FGs (NUP49) to 53 FGs (SONB) and are usually present in the N-terminus of the protein.

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Figure 4.2. Protein localization of MCNC Green fluorescently tagged MCNC (MCNC-GFP) living cells were visualized using the confocal microscopy. Shown here is a projected image of a single germling at four nuclei stage. MCNC-GFP localizes non-uniformly to the cytoplasm of the cell during interphase with no nuclear localization. Arrowheads point to two of the more distinctly visible nuclei. At mitosis, MCNC-GFP is known to re-distribute through out the cell including the nucleus.

109

Figure 4.3. Protein analysis of MCNC using Western blot (A) Protein from endogenously tagged MNCC-GFP was analyzed by Western blot analysis using anti-GFP antibody. MCNC-GFP runs at the molecular size of approximately 200kDa while the predicted protein size for MCNC with GFP is 110kDa. The lower bands in the size range of 105kDa to 75kDa most probably are a result of proteolytic degradation. (B) Protein from exogenously tagged AlcA-MCNC-GFP induced for 0, 1 and 2 hours with threonine were analyzed by Western blot analysis using anti- GFP antibody. AlcA-MCNC-GFP runs at the same size as endogenously tagged MCNC- GFP meaning the size discrepancy is due to protein modification and not due to wrong gene calling.

110

Figure 4.4. Characterization of mcnC deletion The colony size of ΔmcnC was measured at both 32˚C and 42˚C along with appropriate controls. GR5 was used as the wild type (WT) positive control and LPW3 containing the ts nimA1 mutation was used as a negative control. ΔmcnC shows reduce growth at both 32˚C and 42˚C; with a more pronounce effect at 42˚C.

111

Figure 4.5. Bright field and DAPI images of ΔmcnC with wild type To characterize the growth defect of ΔmcnC at 42˚C, spores from both ΔmcnC and wild type (WT) were inoculated in liquid media and grown for 8 hours at 42˚C.(A) shows a wild type germling with 16 nuclei. DIC image shows a polarized germtube while DAPI shows typical stained nuclei. (B) shows ΔmcnC spores with no polarized growth or slow polarization as seen with DIC. DAPI shows the number of nuclei to be anywhere from 4 to 8 in numbers at which point wild type cells always show polarized growth. ΔmcnC thus causes a delay in polarization. These cells however do polarize at later time points.

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Figure 4.6. Effect of alcA-mcnC over expression Wild type GR5 strain transformed with alcA-mcnC was tested for growth on both repressive media and inductive media at 32˚C. alcA-mcnC colonies grew normal and comparable to GR5 under repressive condition but looked sickly under inductive condition.

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Figure 4.7. Effect of over expressed mcnC (A) alcA-mcnC spores germinated in minimal media ethanol at 32˚C for 10 hours (B) alcA-mcnC-GFP spores germinated in minimal media ethanol at 32˚C for 10 hours. Both strains show atypical germlings of varying lengths with multiple branching. Wild type A. nidulans germlings grow in bipolar fashion and subsequent branching occur when cells reach a certain length. Therefore, over expression of mcnC affects polarization.

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Figure 4.8. Effect of mcnC over expression on nucleopore protein SONB Over expression of mcnC by inducible expression of alcA-mcnC in a SONB-GFP tagged strain shows the same multiple germtube phenotype as seen in Figure 4.7. (arrows). Other normal looking cells show SONB-GFP around the nuclear periphery.

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Figure 4.9. mcnC genetically interacts with nimA (A) nimA1 is temperature sensitive at 42˚C. ΔmcnC in a nimA1 background makes it more temperature sensitive than control strains at a lower temperature of 37˚C. Therefore, ΔmcnC shows synthetic lethality with nimA1. (B) nimA7 also shows synthetic lethality with ΔmcnC. Double mutants are sick at 34˚C while the single mutants grow normal. Therefore, mcnC genetically interacts with nimA.

116

CHAPTER 5

MCNB IS A FORK-HEAD DOMAIN CONTAINING PUTATIVE TRANSCRIPTION FACTOR

5.1. Introduction 5.1.1. nimA transcription factors Many forms of control have been shown to regulate the mitotic cell division cycle, such as protein kinase activity, specific proteolytic degradation and changes in intracellular locations. The regulation of gene expression is also shown to play an important role in cell cycle controls. For example, NIMA activity is tightly cell cycle regulated not only post-translationally by multi-stage phosphorylations and proteolysis but also transcriptionally, making its activity largely restricted to a brief window at the G2-M transition and into M (Osmani, May et al. 1987; Pu and Osmani 1995; Osmani and Ye 1996). However, no transcription factors for nimA have been identified to date. There is very little known about how nimA gene expression is rapidly turned on during G2 and turned off in M. Cell cycle regulated transcription has also been studied in the fission yeast, Schizosaccharomyces pombe, and in this regard some light has been shed into the transcriptional control of a group of genes, including, the NIMA related kinase fin1. Fission yeast Fin1 shares many attributes with NIMA. Overexpression of Fin1 induces CDK1 independent premature condensation of chromosomes and Fin1 activity is cell cycle regulated much like NIMA, through cell cycle regulated gene expression and PEST-sequence directed proteolytic degradation. A recent search of the fission yeast genome database using a specific promoter element (PCB) present in cdc15 which confers M and G1 specific transcription, identified the transcription factor complex PBF, and showed it to control the expression of Fin1, and interestingly several other genes

117 involved in mitosis, cytokinesis and mitotic exit (Buck, Ng et al. 2004). In a subsequent genetic screen for cell cycle regulators, two novel fork-head transcription factors Fkh2p and Sep1p were identified and shown to be necessary for PCB-driven M-phase specific transcription. Additionally in budding yeast, the PBF complex was shown to contain the Mcm1 protein along with two fork-head transcription factors, Fkh1p and Fkh2p, which controlled the transcriptional mitotic peak of a cluster of genes important for mitosis, called CLB2 gene cluster. Interestingly, in mutants lacking the two fork-head transcription factors, the genes in the CLB2 gene cluster lost their cell cycle regulation. The altered expression of the CLB2 gene cluster resulted in defects in the structure of the mitotic spindle, nuclear migration, cell morphology and cell separation. Hence, the forkhead transcription factors present in the PCB complex play important roles in the regulation of mitotic gene expression and represent potential candidates that may play a role in transcriptional control of nimA gene expression as well.

5.1.2. Fork-head domain containing transcription factors The forkhead type transcription factors are a class of sequence specific regulators that function in a broad spectrum of cellular and developmental processes in species ranging from yeast to humans. Forkhead transcription factors in eukaryotes have long been implicated in a number of cellular processes including embryogenesis, development, and the cell division cycle. Mutations in genes encoding for forkhead proteins have been linked with several pathological conditions like thyroid agenesis, cleft palate, speech development defects and tumorigenesis. A 110 amino acid DNA binding domain called the forkhead (FKH) /hnf3 domain defines different members of this family. Mutual interactions of various secondary structural elements present in this domain form a three dimensional entity resembling the shape of a butterfly: a core made of α-helical and β- sheet elements flanked by two non-ordered wing like structures. Therefore, this domain is synonymously also called the “winged helix” structure. It represents a unit that cannot be further split without losing its DNA binding properties. In some forkhead proteins, this domain also embeds a nuclear localization signal allowing for its accumulation into the nucleus. Recent studies also reveal the existence of a secondary domain called the 118 forkhead-associated domain (FHA) besides the FKHs for some transcription factor proteins. The FHA domains supposedly act as phosphoprotein binding motifs that participate in regulated protein-protein interactions. Cell cycle dependent phosphorylations of proteins therefore promote interaction with the FHA domain of forkhead transcription factors and may be the underlying reason for cyclical transcriptional changes. In both S. cerevisiae and S. pombe, four forkhead transcription factors are present. Three of the budding yeast genes (FKH1, FKH2, HCM1) participate in regulation of various cell cycle processes whereas the fourth gene (FHL1) codes for a component of the rRNA processing machinery. The Fkh1p and Fkh2p are different from the other two members of the group as they contain the FHA domain. Both these proteins as mentioned before play an important role in the regulation of the CLB genes. The CLB cluster genes are transcribed in late S and G2/M phases and are involved in the G2 to M transition. In fission yeast, two forkhead genes, sep1 and mei4 are involved in the regulation of cell septation and the cytokinetic checkpoint pathway, respectively. The other two genes fhl1 and fkh2 encode proteins with forkhead domains and forkhead associated domains. Fhl1 is highly similar to S. cerevisiae Fhl1p, whereas Fkh2 is more closely related to Fkh1p and Fkh2p of S. cerevisiae. Neither fhl1 nor fkh2 seem to perform essential functions. However, Fkh2 is phosphorylated during the cell division cycle, with a timing that suggests that the posttranslational modification is important for the periodic expression of genes required during M and G1 phases of the cell cycle. Therefore, Fkh2 plays a role in cell cycle dependent regulation of gene expression in S. pombe. A search of the genome database also reveals homolog of all four forkhead transcription factors. Surprisingly enough, a gene identified as a copy number suppressor of the nimA1 mutation turned out to be one of the forkhead transcription factors. All this data suggest that forkhead proteins play conserved roles in regulating cell cycle processes. Studies of metazoans have also linked forkhead proteins with the regulation of cell cycle dependent gene expression. Regulation of forkhead transcription factors regulates expression of mitotic genes like cyclin B and polo-like kinase. Therefore, it is very likely that understanding the mechanisms of regulation of forkhead proteins in the

119 cell cycle of eukaryotes will yield important insights into the regulation of key proteins including NIMA.

5.2. Results 5.2.1. Sequence characterization of mcnB reveals a forkhead domain mcnB encodes for a 78.7kDa protein. The MCNB protein sequence was queried using blastp (Altschul, Madden et al. 1997) against the NCBI genome database to identify homologous proteins from other organisms. As listed in table 5.1, significant homologues of MCNB were identified in all seven Aspergilli species along with homologs in Neurospora crassa, Coccidioides immitis and Saccharomyces pombe. In S. pombe, mcnB is the ortholog of sep1.A domain with significant sequence homology to the forkhead domain found in other transcription factors (Fkh2 from S. cerevisiae and Fkh2 from S. pombe) was identified in MCNB and matched to a stretch of 110 amino acids from aa 206-315. See Figure 5.1. No FHA domain however was observed. Additionally, the pestfind software at http://emb1.bcc.univie.ac.at/embnet/tools/bio/PESTfind/ found a number of PEST sequences, the majority of which were localized to the C terminus of the protein. PEST sequences in many proteins are known to play a major role in ubiquitin mediated protein degradation (Rechsteiner and Rogers 1996).

5.2.2. MCNB localizes to the nucleus during G2-M transition mcnB was endogenously tagged with green fluorescent protein at its C-terminus using the fusion PCR technique. The tagged strains were confirmed by diagnostic PCR and Western blot analysis (Figure 5.2A).MCNB-GFP cells were then visualized using live cell microscopy. As can be noticed from figure 5.2B, one can see MCNB-GFP to localize in the nucleus of many cells. However, occasionally there were cells with no localized MCNB-GFP signal. Moreover, the average intensity of signal in the nucleus varied from one cell to another. This suggested that the levels of MCNB may be regulated through the cell cycle and localized to the nucleus at particular cell cycle stages. In order to test this, MCNB-GFP tagged strains were crossed to a strain containing

120 histone H1 tagged with red fluorescent protein. This allowed the visualization of MCNB in relationship to the DNA. Spores from the double tagged strain were then germinated and visualized for both green and red fluorescence under the confocal microscope. As once can see from figure 5.3, MCNB-GFP proteins begins to accumulate in the nucleus during G2 (panel a, b) and reaching its peak just prior to mitotic entry (panel c, d). As DNA begins to condense and prepare for entry into M (panel e), MCNB disappears from the nucleus. The DNA continues to condense and undergo nuclear segregation and enters G1 (panel f). If these cells are followed for another round of cell cycle, the re- accumulation of MCNB can be observed in the next G2 and the process repeated all over again (panel g, h).

5.2.3. mcnB is a non-essential gene The mcnB gene was replaced with the A. fumigatus pyroA nutritional marker using homologous recombination in the wild type strain GR5. The null allele for mcnB was confirmed using diagnostic PCR and analyzed for heterokaryon formation to assess for mcnB essentiality. ΔmcnB strains were viable and did not form any heterokaryons. Additional growth tests were done on a variety of growth conditions as mentioned in section 2.11 to check for potential phenotypes. No significant phenotypic changes were noticed in mcnB deleted strains.

5.3. Discussion 5.3.1. mcnB up-regulation of NIMA1 is likely due to increased transcription The over expression of mcnB in nimA1 cells result in the up-regulation of the NIMA1 protein levels (Figure 3.7). It is plausible therefore that the increased levels of NIMA1 protein compensates for the low activity of NIMA1 protein kinase activity in nimA1 strains thereby leading to a suppression of the G2 arrest. The presence of a forkhead domain, commonly found in other transcription factors strongly supports this idea. However whether MCNB directly activates nimA transcription or through the activation of some other gene which in turn affects NIMA protein levels is not yet known. Moreover, mcnB has strong homology to the S. pombe sep1 gene. Sep1p, a

121 forkhead containing transcription factor, is thought to play an important role during PCB driven M phase transcription of a number of genes with roles in regulating sister chromatid separation, septation and cytokinesis. Expression of Sep1p, along with a second forkhead protein Fkh2p, results in significant changes to the mRNA levels of the fin1 gene and is required for the periodic accumulation of Fin1p during M (Buck, Ng et al. 2004). Fin1 is considered the nimA related kinase in S. pombe. Therefore, a role for forkhead transcription factor expression in regulating a nimA like kinase exists in the fission yeast and suggests that a similar regulation may exist in for nimA kinase. Additionally, the forkhead transcription factors from budding yeast, Fkh1p and Fkh2p, regulate the CLB2 gene cluster at S and G2 to M transition, a lack of which results in aberrant mitosis. Similarly, the fhl1 and fkh2 gene from S. pombe regulate mitotic entry by influencing gene expression of G2 and M specific proteins.

5.3.2. MCNB and NIMA show identical protein expression profiles MCNB-GFP protein localizes to the nucleus in G2 and continues to accumulate in the nucleus reaching a maximum level at M. At mitotic entry, the nuclear retention of MCNB-GFP is lost and the protein is either degraded or dispersed into the cytoplasm. The rapid kinetics of the loss of MCNB signal suggests a rapid degradation of the protein and the presence of multiple PEST sequences in the protein support this idea. NIMA protein levels also rise during late G2, peaking during M and get degraded at mitotic exit. The pattern of protein expression of MCNB is thus identical to that of NIMA but its initiation and degradation seems to precede that of NIMA, which is consistent with it being a potential transcription factor for nimA.

5.3.3. Functional redundancy of forkhead transcription factors In both S. cerevisiae and S. pombe there are four forkhead transcription factors, all known to have specific cell cycle regulatory functions but none considered to be essential. The homologs for all four forkhead genes are also found in A. nidulans (refer table?). Therefore, mcnB deletion not resulting in a lethal phenotype suggests a functional redundancy shared among the four transcription factor proteins. In the recent past, NIMA

122 functions have been implicated in many other cell cycle processes and cellular locations as mentioned in chapter 1. The presence of four transcription factors with overlapping functions may be required to efficiently regulate these aspects of the NIMA protein. Thus, among the various mechanisms of regulating NIMA kinase activity, which include its regulation through sub-cellular localization (NIMA localizes to the cytoplasm in G2 and enter the nucleus only at M), regulation of NIMA abundance ( NIMA protein levels begin to accumulate in G2, peak at M and get degraded at exit), regulation by degradation (NIMA protein and mRNA levels are rapidly degraded at mitotic exit which for the protein involves the APC) and post translational cell cycle specific regulation by phosphorylation, MCNB may add to a new mode of regulation through transcriptional control. By activating and limiting the function of MCNB to a window of G2 to M, mcnB may serve to help control nimA mRNA levels during the cell cycle.

123

A

mcnB (length 717) Table 5.1. mcnB homologs Organism Length Identity E value % mcnB encodes for a 717 amino acid protein. Aspergillus 725 66 0.0 (A)Blastp homology searches using mcnB fumigatus Aspergillus 722 66 0.0 protein sequence identified proteins sharing oryzae significant homology in all members of the Aspergillus 604 69 0.0 Aspergilli family and in fungus Coccidioides clavatus Neosartorya 722 66 0.0 immitis, Neurospora crassa, and fischeri Schizosaccharomyces pombe and Ustilago Aspergillus 717 63 0.0 flavus maydis. (B) Blastp of the Aspergillus genome Aspergillus 717 63 0.0 database using mcnB protein sequence also niger finds 3 additional proteins with homology to Aspergillus 723 67 0.0 terreus MCNB, all with a conserved forkhead domain. Coccidioides 724 52 3e-131 Similar orthologs are found in both immitis Neurospora 805 36 5e-53 S. cerevisiae and S. pombe. crassa S. pombe 663 54 5e-29

B

mcnB (length 717) Organism Length Identity E value % Aspergillus 711 47 3.6e-23 nidulans Aspergillus 1095 28 6.5e-12 nidulans Aspergillus 634 27 2.7e-10 nidulans

124

MSPKQPAKRVLGAHSSQDRTTTIKPQPRLEPSPLPLQSTENPPSHHKYVLNPTSTSP NRVSPTKFSRESRTSGCAQARLGYVPITAPAPPTFTTDSPIKKIPLETYSHPTPSSA PMPQSALFTTFSSVRNVKHTTTGDLDVDVPSADTFADFPEPSQLAKHSFKRSLLDAA PLKERTKKPKGEEVTTVQLPEPHELPPIEDDGTKPPYSYATLIGMSILRAPNRRLTL AQIYRWISDTFSYYKNSDPGWQNSIRHNLSLNKAFIKQERPKDDPGKGNYWAIEPGM ETQFIKDKPVRRATMTSMPTLSITPQQEPTYSQGSSATTWAVPPPAQHPVSKSSKHV DLSSDATIPASDPALQEDIGDDTAACLTTNPPRSSPPQPIHSSPPVAPPRFARPATP PTPCHPSIPSDGPRHRKRKSNTMNDSGYFSSLESSAMRSNKASHYLTSDTDIEPPRI KRGRAEEEIARIRSSSRDISPSHSGYLKETGIVVGSSPVRNEYINVLAGPLTPVIKF KKPAKPPPSVSPNTNLRNHRKKIQHMVNSPIKRLGLDEDLPWSPAFNIHDEAYTPHD GLHVSFDVFADPTTEPVSNPAYGSPEKRSAKRARTEAHGPTGNALADITALSANNRI GGLKPLSPNKSKRLFFSDSPSKLPDSGRFIDSAHDDFFSWHLFDDSPQEVDGVDLLQ GFQKIGGSSKDDASKSRSHISQPTFQRGSNTRP#

Figure 5.1. MCNB protein sequence mcnB encodes for a 717 amino acid protein with a molecular weight of 78.7kDa. The protein contains a conserved forkhead domain found in known transcription factors shown here in blue and contains many PEST sequences known to target proteins for degradation by the ubiquitinated mediated APC system.

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Figure 5.2. MCNB protein analysis and localization (A) Green fluorescently tagged MCNB (MCNC-GFP) were confirmed by western blot analysis. Multiple lower bands are most probably proteolytic degradation products. (B) Live cells were visualized using the confocal microscopy. Shown here is a projected image of a few germling at 1-2 nuclei stage. MCNB-GFP localizes to the nuclei only in some germlings and not all. Moreover, the signal intensity in nuclear accumulated cells vary suggesting cell cycle regulated localization of MCNB.

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Figure 5.3. MCNB localization through the cell cycle MCNB-GFP proteins begins to accumulate in the nucleus during G2 (panel a, b) and reaching its peak just prior to mitotic entry (panel c, d). As DNA begins to condense and prepare for entry into M (panel e), MCNB disappears from the nucleus. The DNA continues to condense and undergo nuclear segregation and enters G1 (panel f). If these cells are followed for another round of cell cycle, the re-accumulation of MCNB can be observed in the next G2 and the process repeated all over again (panel g, h).

127

CHAPTER 6

MCNA IS A NOVEL SUB-NUCLEAR PHOSPHOPROTEIN WHICH TRANSIENTLY LOCATES TO CYTOPLASMIC BODIES SPECIFICALLY DURING MITOSIS AND G1 OF THE CELL CYCLE

6.1. Introduction 6.1.1. Regulation of gene function through protein/mRNA turnover Initiation of mitosis in Aspergillus nidulans requires the cooperative action of two cell cycle regulated protein kinases, CDK1/cyclin B and NIMA which both have peak activity during mitosis. Transcription plays a major role in regulating the mRNA levels of nimA (Osmani, May et al. 1987). In addition, both kinases are inactivated during mitotic progression and this cell cycle specific inactivation is required for proper mitotic exit. This critical step involves ubiquitin mediated degradation of cyclin B and NIMA proteins by the anaphase promoting complex, APC (Osmani and Ye 1996; Ye, Fincher et al. 1998). The mRNA levels of nimA also catastrophically collapse during exit from mitosis but much less is known about the mechanisms underlying this regulation. However, degradation of the cyclin B and NIMA transcripts would be expected to be essential for tight control of their mRNA levels. Of the three copy number suppressors identified for nimA1, mcnA results in up- regulation of NIMA1 protein when over expressed, much like mcnB (figure 3.7). However, MCNA protein does not contain any distinctive motif/domain suggestive of its role as a transcriptional regulator of nimA. Instead, as will be shown in this chapter, MCNA locates to the nucleolus predominantly and changes it localization to discrete bodies in the cytoplasm during specific phases of the cell cycle. We hypothesise that the

128 specific localization pattern of MCNA may play a potential role in regulating NIMA levels through the cell cycle through post-transcriptional or post-translational modulation.

6.1.2. Regulation of protein activity by nucleolar sequestration Cell cycle regulation of protein localization to specific sub-cellular compartments, like the nucleolus, is one of the many effective ways to regulate specific activities during the cell cycle. For example, in budding yeast, the protein phosphatase Cdc14 is a key regulator of late mitotic events enabing cells to exit mitosis through dephosphorylation of key proteins involved in exit from mitosis. Cdc14 levels are constant through the cell cycle throughout G1, S/G2 and early mitosis. However, Cdc14 is kept inactive due to its sequesteration in the nucleolus by association with specific inhibitory proteins. The phosphorylation of Cdc14 and associated proteins in late anaphase allows the release of Cdc14 from the nucleolus so that it might reach its targets in the nucleus and the cytoplasm (Stegmeier and Amon 2004). Cdc14 is then re-sequestered in the nucleolus during late telophase. Thus the restriction of Cdc14 to the nucleolus during a specific window of the cell cycle allows the nucleolus to serve a secondary role as a sequesteration center to help regulate passage through the cell cycle. Like Cdc14, potentially other proteins that localize to the nucleolus may not act solely (or even primarily) in the nucleolus but may be accumulated there in anticipation of their eventual release at specific cell cycle stages ensuring regulated activity towards targets entities.

6.1.3. Programmed degradation of mRNA take place in P bodies Eukaryotic mRNAs are degraded by several mechanisms (Beelman and Parker 1995). One major mRNA degradation mechanism involves initiation of mRNA decay by shortening of the 3’ poly(adenosine) tail, followed by removal of the 5’cap by decapping enzymes which in turn allows 5’ to 3’ exonucleolytic decay (Sheth and Parker 2003). In yeast, mRNA decapping and 5’ to 3’ degradation occur in discrete cytoplasmic foci called processing bodies (P bodies) (Sheth and Parker 2003). Enzymes required for RNA decay and the resulting RNA degradation intermediates localize to P bodies and dynamic changes to their sizes occur depending on the global state of RNA turnover. The decrease

129 in P bodies when deadenylation is inhibited, and the increase in P bodies when decapping or 5’ to 3’ exonucleolytic digestion is inhibited, is suggestive of P bodies being specific sites of mRNA decay. In addition, treatment of cells with cycloheximide, which rapidly inhibits decapping by trapping mRNA on polysomes, results in complete loss of P bodies suggesting that P bodies are sites of mRNA decapping. The occurrence of RNA decay in P bodies thus add to a growing number of specific sub-cytoplasmic compartments that are known to sequester mRNA molecules (Anderson and Kedersha 2002) and are known to play a role in mRNA metabolism. Moreover, similar cytoplasmic foci containing specific P body proteins have been observed in mammalian cells (van Dijk, Cougot et al. 2002). Recent data has also established the presence of RISC-mRNA complexes in P bodies which are the main players helping mediate RNAi mediated post-transcriptional gene silencing (Rossi 2005). Thus, P bodies represent key elements in the regulation of controlled mRNA degradation and thus protein expression.

6.1.4. Cyclin B mRNA is degraded during mitosis via a specialized P body Direct evidence for mRNA degradation promoting cell cycle progression has come from the study of the RNase mitochondrial processing (RNase MRP) protein. RNase MRP is a site specific endonuclease ribonucleoprotein (Chang and Clayton 1987). The protein and RNA component of RNase MRP are highly conserved in structure and sequence in all eukaryotes (Lygerou, Mitchell et al. 1994; Jarrous, Eder et al. 1999; Cai, Aulds et al. 2002). Although originally isolated from mammalian mitochondria, the majority of RNase MRP localizes to the nucleolus (Reimer, Raska et al. 1988) and plays a role in rRNA processing (Chu, Archer et al. 1994; Henry, Wood et al. 1994). In both budding and fission yeasts, mutation in the MRP RNA causes cell cycle delay (Paluh and Clayton 1996; Cai, Aulds et al. 2002) while RNase MRP mutants shown genetic interactions with S. cerevisiae cell cycle mutants that have mitotic exit functions (Gill, Cai et al. 2004). RNase MRP mutants accumulate late in mitosis and exhibit characteristics similar to “exit from mitosis” mutants (Surana, Amon et al. 1993). Western and northern analysis show that cyclin B protein and mRNA accumulate in such mutants and that the increased CDK1/cyclin B activity causes the mutant phenotype (Cai,

130 Aulds et al. 2002). Further studies implicate RNase MRP in the cleavage of the 5’ untranslated region of cyclin B mRNA both in vitro and in vivo (Gill, Cai et al. 2004) resulting in rapid degradation and regulation of cyclin B levels. Note that RNase MRP must be localized to the nucleolus through the cell cycle to process ribosomal RNAs while degradation of cyclin B is presumed to occur in the cytoplasm. Therefore the activity of RNase MRP against cyclin B is regulated during the cell cycle by changing its intracellular localization (Gill, Aulds et al. 2006). During metaphase nucleolar RNase MRP re-localizes to the entire nucleus and to a single cytoplasmic focus that persists till telophase. After nuclei complete division, RNase MRP moves back into the nucleolus. The localization of RNase MRP to the cytoplasmic foci is a temporally and spatially regulated event. It appears only in daughter cells during mitosis and is rarely visualized in the parent cell. Because the focus appears in a cell cycle controlled manner, and only in the daughter cell, the RNase MRP is said to constitute a temporal asymmetric MRP body, or TAM body (Gill, Aulds et al. 2006). Since cyclin B mRNA needs to be degraded only during a certain time of the cell cycle, the change in RNase MRP localization via the TAM body presents a simple but efficient means to regulate the activity of RNase MRP towards cytoplasmic cyclin B during mitotic exit. The TAM body can also be considered to be a specialized P body as it seems to contain the same protein component (Xrn1p) required for mRNA degradation as P bodies and serves an identical function, mRNA decay.

6.2. Results 6.2.1. mcnA sequence reveals a novel protein specific to the Aspergilli The protein sequence of MCNA shows several identifiable domains when searched with the motif resource software ELM (http://elm.eu.org/). MCNA, a protein of 506 amino acids (Figure 6.1), contains a globular domain (2-270 aa), a coiled-coil domain (120-150 aa), four APC degradation-like box motif with RXXL sequence (88-93 aa, 167-172 aa, 187-192 and 425-430 aa), substrate recognition site that interacts with cyclin thereby increasing phosphorylation by CDK/cyclin (RLLEL sequence, 51-55 aa) and seven potential CDK1 phosphorylable consensus sites. The protein sequence of

131 mcnA was queried at the NCBI database using blastp (Altschul, Madden et al. 1997) to identify putative homologous proteins from other organisms. Proteins sharing significant homology were identified only in the seven Aspergilli species and the pathogenic fungus Coccidioides immitis (Table 6.1). MCNA therefore represents a novel protein highly conserved among the Aspergillus species which is also present in C. immitis.

6.2.2. MCNA sub-cellular localization 6.2.2.1. MCNA localizes to a single focus in the nucleus The mcnA gene was endogenously tagged with green fluorescent protein using the fusion PCR protocol to study its protein localization. A MCNA targeted GFP fusion construct was used to transform wild type GR5 strain and successful transformants chosen by prescreening for GFP fluorescence. In figure 6.2A, a single fluorescent focus (hereby termed as a MCNA body) per spore was observed. The positive transformants were then confirmed for proper tagging by performing diagnostic PCR and the protein size confirmed using western blot analysis. MCNA-GFP is predicted to be 56.6 kDa but appears to migrate as several specific molecular species (Figure 6.2B). When GFP tagged cells are grown, fixed and stained with DAPI, a single MCNA body is observed within the nuclei in about 90% of the cells (Figure 6.2C). In the remaining 10% of the cells, the MCNA body localize to the cytoplasm, sometimes as more than single bodies.

6.2.2.2. The MCNA body is located near the nuclear envelope When the ER is visualized using a red fluorescent tagged ER marker Erg4, MCNA-GFP localizes near the nuclear periphery (Figure 6.2D). Note that in DAPI stained cells, the MCNA body localizes to the dark shadow of the nucleus, which represents the nucleolus that remains unstained by DAPI. The nucleolus is always in close association with the nuclear envelope as well and therefore this led to speculation that MCNA may be localized to the nuclear periphery, near the NE due to its association with the nucleolus. I therefore performed co-localization studies of MCNA with the nucleolus.

132 The ANA1 antibody is an autoimmune antibody that recognizes the human fibrillarin nucleolar protein and can be used as a marker for the nucleolus during immunofluorescence studies (Kuhn, Jarzabek-Chorzelska et al. 1990). When MCNA- GFP tagged cells are fixed and stained for ANA1, one can visualize the MCNA dot near the nucleolus (Figure 6.3A). However, the area occupied by MCNA is only a sub-domain of the nucleolus as shown in figure 6.3B and locates at the periphery and not to the interior of the nucleolus. To confirm the immunofluorescence data, we doubly tagged wild type strains for MCNA and a number of nucleolar proteins. As can be seen in figure 6.3B, three different nucleolar proteins, Fibrillarin, Bop1 and CgrA, endogenously tagged with red fluorescence protein, when visualized along with MCNA-GFP, clearly showed MCNA to localize to a sub-domain of the nucleolus.

6.2.3. MCNA location varies during mitosis and G1 of the cell cycle Live imaging using confocal microscopy was performed to study the localization of MCNA during the cell cycle. Spores containing MCNA-GFP were inoculated in a microdish and incubated at room temperature to allow growth of germlings to the 4-8 nuclear stage. Each germling contained 4-8 MCNA bodies, one per nucleus, each of which were then followed through the cell cycle. The MCNA bodies were observed to double with the number of nuclei during a single cell cycle and undergo a unique pattern of segregation during and after mitosis (Figure 6.4). In order to visualize the relationship between MCNA and DNA, the MCNA-GFP tagged stain was crossed to a RFP tagged histone H1 strain, to generate a doubly tagged strain. The strain shows the exact same pattern of segregation for MCNA. The beginning of M phase is marked by the condensation of DNA (figure 6.4, panel a) followed by DNA segregation into two daughter nuclei (panel b). During DNA condensation, a single MCNA body remains associated with the nucleus but as the DNA separates (panel a), the MCNA body is excluded from the nucleus and appears in the cytoplasm (panel b, arrowhead). About 10 mins after completion of mitosis MCNA protein begins to appear in the daughter nuclei (panel c). The single cytoplasmic MCNA body at this time also divides into a slightly variable number of bodies, usually 2-4 (panel d), which linger in the cytoplasm for a

133 period of time (panel e). It is only after ~35 mins of its first appearance in the daughter nuclei that the multiple cytoplasmic MCNA bodies completely disappear from the cytoplasm (panel f). The cell cycle stage at this point of time coincides with the cell being past G1 entering S-phase. Therefore, MCNA is a nuclear protein localized in close vicinity of the nucleolus during interphase but is cytoplasmic in late mitosis and through most of G1. Noticeably MCNA divides independently of DNA segregation and completes its division during G1.

6.2.4. MCNA co-localizes near nucleolar proteins in interphase but for only part of mitosis As mentioned before, MCNA GFP co-localizes to a sub-domain of the nucleolus when visualized with three different nucleolar proteins (Fibrillarin, Bop1 and CgrA). We then visualized localization of MCNA-GFP in comparison with red fluorescent tagged nucleolar proteins through the cell cycle. During interphase, MCNA-GFP localized as a dot to a sub-domain of the nucleolus as described above. When chromosomal segregation occurred, both MCNA-GFP and nucleolar proteins were localized to the cytoplasm and still co-localized. However, within 1-2 minutes, MCNA splits into a number of smaller bodies not associated with the nucleolar proteins. The nucleolar proteins begin to reappear in the daughter nuclei before MCNA and complete their reassembly into the daughter nuclei before MCNA is completely removed from the cytoplasm. Hence, even though there is a over lap of localization between MCNA and nucleolar proteins, MCNA does not behave similarly to nucleolar proteins during mitosis and its segregation pattern and timing is unique from the nucleolus occurring during G1.

6.2.5. S-tag affinity purification of MCNA A good approach to further study the function of any protein is to identify interacting partners of the protein. This can be achieved directly by performing an affinity purification of the protein of interest. Therefore, we constructed an S-tag targeting construct that could be used to tag mcnA and so purify it along with its potential binding partners. The S-tag peptide, due to its small size (15aa), provides minimum interference

134 to the tagged protein and possesses high affinity to S protein. The S-Tagged chimeras can therefore be purified using beads crosslinked to the S-protein. An S-tagged mcnA strain was constructed by transforming ΔKu70 strain SO451 with a targeted S-tag cassette. Successful transformants were confirmed by diagnostic PCR and western blot analysis (refer protocol 2.5.2). Total protein extracts were then used for affinity purification with S-protein beads. Purified MCNA-S-tag was eluted from the beads using SDS protein sample buffer and run on SDS-PAGE. The gel was stained with Coomassie blue to visualize the purified proteins. Figure 6.2B shows the different protein bands from an affinity purified MCNA sample. A control sample similarly purified from protein extracts from a strain not containing a tagged protein was also performed. A parallel western blot analysis using 1/100th of the total protein used for the purification was performed using an anti-S-tag antibody which confirmed the 4 different purified bands to be MCNA. Therefore, MCNA migrates as several specific molecular weight species but does not co- purify with any high abundance interacting partners.

6.2.6. MCNA is a phosphoprotein When visualized with western blot analysis using anti-GFP antibody or anti-S-tag antibody the tagged versions of MCNA appears to migrate as several specific molecular species, suggesting potential post-translational modification to the protein. As mentioned before, seven CDK1 specific phosphorylation sites are found in MCNA and therefore provide the possibility of it being modified through cell cycle specific phosphorylation. Many proteins important for cell cycle progression are regulated during the cell cycle through phosphorylation events and the phosphorylation of MCNA may be important for its function. In order to confirm that phosphorylation of MCNA is responsible for its appearance as multiple bands, protein phosphatase experiments were performed. Protein phosphatases specifically target phosphate groups on a protein and cleave them, essentially de-phosphorylating a phosphorylated protein. In order to perform the experiment, beads containing affinity purified MCNA were divided equally into three tubes (A, B and C).Tube A was equilibrated with a phosphatase buffer, tube B was equilibrated with phosphatase buffer containing a protein phosphatase. To tube C, the

135 protein phosphatase was added but after equilibrating it with a phosphatase inhibitor buffer. The inhibitor buffer inactivates the protein phosphatase enzyme. Eluted protein from each tube was then run on SDS PAGE and the gel silver stained after protein separation. Figure 6.5A shows the gel with all three lanes. Note lane A, shows the 4 different bands of MCNA. Lane B is protein phosphatase treated and all the bands collapse to a single size that corresponds to the predicted 56.6 kDa of full length MCNA. Lane C on the other hand looks like lane A proving that the collapse of bands in lane B is due to the specific activity of the protein phosphatase removing the phosphate groups from MCNA. Therefore, MCNA is a phosphoprotein, likely phosphorylated on multiple sites.

6.2.7. Three potential CDK1 phosphorylation sites of MCNA identified by Mass spectrometric analysis Coomassie stained, affinity purified MCNA-S protein was identified by capillary- liquid chromatography-nanospray tandem mass spectrometry (Nano LC/MS/MS). In this technique, the protein of interest is proteolytically digested and the resulting peptides chromatographically separated and analyzed using tandem mass spectrophotometry. Using this method, 56% sequence coverage was achieved (Figure 6.5B) which covered three of the seven potential CDK1 specific phosphorylation sites. Modification by phosphorylation was detected in four peptide fragments and the phosphorylation confirmed to be at Serine 330, Serine 365 and Serine 442 (Figure 6.5B, highlighted in blue). All three phosphorylated sites conform to the known minimal CDK1 consensus site “SP” and therefore suggest a potential cell cycle regulation of MCNA by CDK1 mediated phosphorylation.

6.2.8. mcnA deletion and mutation of MCNA phosphorylation sites A null mcnA allele was generated using homologous integration to replace its coding region with the A. fumigatus pyroA nutritional marker. The deletion construct was generated using fusion PCR (Yang, Ukil et al. 2004) and transformed into the wild type GR5 strain. Homologous integrants were confirmed by diagnostic PCR. Deleted

136 transformants were checked for heterokaryon formation to assess for mcnA essentiality. mcnA deletion (ΔmcnA) did not lead to heterokaryon formation and hence mcnA is not an essential gene. Further characterization of ΔmcnA was performed by testing under an array of growth conditions as mentioned in section 2.11 to uncover potential conditional phenotypes. ΔmcnA showed no visible mutant phenotypes and thus mcnA is non- essential. As mentioned before, MCNA is a phosphoprotein and at least three phosphorylable sites were identified by the mass spectrophotometry analysis. We considered that the phosphorylation of MCNA may be essential for its normal function. For instance, a mutant version of MCNA unable to under go phosphorylation at all three mapped phosphorylation sites may prevent, or modify, the normal cell cycle segregation of MCNA. To address the in vivo roles of phosphorylation of MCNA, a construct with three serine-to-alanine point mutations targeted at all three phosphorylated serine sites was generated (mcnA-3A). The construct was designed such that it could transform a strain and result in the gene replacement of the endogenous wild type copy of mcnA with the mutated mcnA3A version. However, in order to perform this replacement successfully I needed to make sure that during transformation mcn3A does not under go any recombination at the endogenous mcnA resulting in a gene conversion. Therefore, a ΔmcnA strain was used such that the replacement construct would replace the marker at the deleted mcnA locus with no possibility of homologous recombination causing gene conversion. The mcnA-GFP-pyrGAF sequence was PCR amplified from the endogenously tagged MCNA-GFP strain and cloned into the TOPO1 vector. Three rounds of consecutive site-directed mutagenesis was performed to generate the mutant insert “MCN3A” in which the serines at positions (330, 365, 442) were converted into non- phosphorylable alanines. The mutant insert was then cut out from the vector by restriction enzyme digestion and used to transform the ΔmcnA strain. ΔmcnA in which mcnA is deleted with the fumigatus pyroAAF marker also contains the pyrG89 mutation in the background allowing for the gene replacement of pyroA with the MCNA3A mutant insert

137 without the possibility of undergoing a gene conversion (MCNA3A converting to wild type MCNA) due to a lack of any homologous MCNA sequence. Transformants that underwent recombination with the mutant MCNA3A insert now grew in media lacking uridine and uracil as they contained the pyrGAF marker. Secondarily, if the transformants were a result of homologous recombination and the cassette integrated at the mcnA locus replacing the pyroAAF marker, they became sensitive to lack of pyridoxine. Hence a double selection strategy for pyrG positive and pyroA negative was used to select the required transformants. Western blot analysis was performed to check if targeting the three serine sites as mentioned above resulted in the complete abolishment of phosphorylation for MCNA. As can be seen from figure 6.6A, MCNA3A still migrated as two specific bands suggesting the existence of other potential phosphorylation sites not targeted by the site directed mutagenesis. The partially phosphorylable MCNA3A strain was tested on an array of growth conditions to check for possible conditional mutant phenotypes. MCNA3A exhibited no growth defects and had normal looking DAPI stained nuclei. However, live cell microscopy of GFP-tagged MCNA3A mutant strains revealed differences in the amount of MCNA per nucleus indicative of a segregation defect for MCNA3A-GFP during mitosis. When visualized through a cell cycle, the cytoplasmic MCNA3A-GFP foci during mitotic segregation appear to be similar in numbers but persist in the cytoplasm for a longer period of time. The time taken to complete the process of segregation of MCNA3A-GFP dots during mitosis and into interphase was thus longer than the time taken for segregation of wild type MCNA-GFP. Another noticeable defect noticed was the unequal segregation of the mutant MCNA3A-GFP (Figure 6.6B). Wildtype MCNA- GFP segregates to each nucleus within a germling in a very even manner such that all nuclei receive an equal amount of MNCA at the end of the segregation process. However, the amount of MCNA3A per nucleus was markedly different when compared to the equal distribution of the wild type protein. Western blot analysis shows an up-regulation of MCNA-3A protein in the mutant strain and this may be a cause for the unequal amounts (Figure 6.6C). The unequal segregation of MCNA3A to the two daughter nuclei and the prolonged persistence of MCNA3A foci in the cytoplasm during its segregation thus

138 strongly suggest that phosphorylation of MCNA plays an important cell cycle role in its segregation.

6.3. Discussion 6.3.1. MCNA protein localization is cell cycle regulated The localization of the MCNA body is very specific during the cell cycle. The MCNA body is intra-nuclear in close proximity to the NE and associated with the nucleolus through S/G2 and most of M. In late anaphase, the MCNA body changes its localization from the nucleolus to the cytoplasm. The MCNA body appears in the cytoplasm only after chromosomal segregation and remains in the cytoplasm beyond mitotic exit forming distinctive numbers of smaller bodies. At the same time MCNA also begins to re-accumulate in the two daughter nuclei. The cytoplasmic bodies, though smaller in size, however persist in the cytoplasm till late G1, before being equally re- distributed to the daughter nucleoli. Hence the localization of MCNA to the cytoplasm is very specific to a late M-G1 window of the cell cycle demonstrating MCNA to be cell cycle regulated. During mitotic exit and G1 entry of the cell cycle, a rapid down regulation of key proteins like cyclin B and NIMA is known to take place. The specific change in sub-cellular localization of MCNA coincides with this M to G1 transition. A rudimentary hypothesis is that specific localization of MCNA through the cell cycle plays a role in NIMA protein and/or mRNA turnover, such that its association with the nucleolus during mitosis may help stabilise NIMA while its cytoplasmic presence during late M /G1 may be essential to regulate NIMA for mitotic exit.

6.3.2. MCNA bodies have some similar features to TAM bodies There is some resemblance between the mitotic dynamics of MCNA to the recently identified TAM body in S. cerevisiae. RNase mitochondrial RNA processing (MRP) proteins, as mentioned before, predominantly localize to the nucleolus, however after mitotic initiation it localizes through out the cytoplasm and to a distinct cytoplasmic foci, the TAM body. The TAM body is in the cytoplasm of budding yeast during metaphase and asymmetrically localizes to the daughter cells. Localization of the TAM

139 body at the specific cell cycle stage is thought to allow for degradation of daughter cell- localized mRNAs, such as cyclin B and regulate mitotic exit. Its cytoplasmic localization also persists until the completion of telophase. Thus there are some basic similarities between the TAM and MCNA bodies.

6.3.3. MCNA is cell cycle regulated by phosphorylation MCNA is a phosphorylated protein and protein made from asynchronously growing cells migrates as several molecular weight species. Using mass spectrometric analysis, at least three potential CDK1 phosphorylation sites were identified in MCNA. A triple mutant version of MCNA protein, MCNA-3A, in which three of the serine sites are mutated still migrated as two molecular weight species. In vitro protein phosphatase treatment show the extra bands to be phosphorylation specific providing evidence of additional sites being phosphorylated in MCNA. The partially phosphorylable MCNA-3A protein shows a segregation defect for MCNA during G1. The MCNA body localizes to the cytoplasm at telophase and stays in the cytoplasm during G1 similar to wild type MCNA. The duration of the MCNA bodies in the cytoplasm is however longer than wild type and also results in an unequal distribution in the daughter cells in late G1. Since the inhibition of phosphorylation of MCNA causes a specific defect during G1, it suggests that phosphorylation plays a role in cell cycle regulation of MCNA. In addition, phosphorylation of MCNA may change through the cell cycle (quantity and/or specific sites) and if mcnA regulates nimA, this would add to a novel mechanism of cell cycle regulation of the M-G1 transition through spatial and temporal control of the regulator itself.

6.3.4. MCNA over expression leads to NIMA up-regulation: a contradiction? The hypothesis that MCNA may function in NIMA regulation during mitotic exit would assume that over expression of MCNA will lead to a down regulation of proteins like NIMA. Rather, MCNA over expression leads to NIMA up-regulation (Figure 3.7). How can this contractidory result be explained?

140 Even though affinity purification of MCNA yields no other interacting proteins, it is highly unlikely for a specific cell cycle regulated structure like the MCNA body to contain only MCNA protein. If MCNA is involved in protein turnover of NIMA, it is possible that MCNA may be in a complex with other proteins that together have a cell cycle function in regulating NIMA. If the proteins in this complex exist in a balanced stoichiometric ratio, over expression of just one protein (in this case MCNA) may lead to inactivation of the complex. Such an example is observed in the case of the dynactin complex, where over expression of one dynactin subunit results in the collapse of the entire complex and its subsequent inactivation preventing its accessory role in cytoplasmic dynein function (Burkhardt, Echeverri et al. 1997). Thus it is likely that MCNA is present in a complex with other proteins in the cytoplasm and this interaction of MCNA with other proteins may be sensitive to the purification protocol used leading to an inability to affinity purify interacting proteins. Thus, the over expression of MCNA may lead to the inactivation of a complex that may have a function in NIMA degradation and thereby lead to up-regulation of NIMA protein causing suppression of nimA1.

141 mcnA (length 511) Organism Length Identity % E value Aspergillus fumigatus 503 74 0.0 Aspergillus oryzae 413 73 0.0 Aspergillus clavatus 503 75 0.0 Neosartorya fischeri 503 74 0.0 Aspergillus flavus 506 72 0.0 Aspergillus niger 504 73 0.0 Aspergillus terreus 516 72 0.0 Coccidioides immitis 480 47 2e-106

Table 6.1. Homologs of mcnA mcnA encodes for a 511 amino acid containing protein. Blastp homology searches using mcnA identified proteins sharing significant homology in all members of the Aspergilli family with similar protein sizes and in the fungus Coccidioides immitis.

1 MTATLERSLSRTSSVSMPLSPMISFRHEATTPISESSLSHIHERLSAIES 51 RLLELRSTALTKDGYVDRRNREDEHIRREFEAHRSISNRIDLNVVALRAD 101 VGQLKSGILQLKSSLGQAGNETVFLRSDVDRLSKNVDQIQVDLEHMQTDV 151 CGCRVEISKLQSAISQLRTELITLQHETSRHLNSFLERFSLIEARMKHSE 201 RVRFNSLAHTTHAPITPVPIVEEDGSLRWPDYFPRTVWRFWCLKKRSRHN

251 RLAQLAEFYQLGGYEYWGRMHQTDGMFSDSDSSDSSDCPSNLTRAEAVRM

301 FPEAAHQALAATLGLVYYKIRNEVGDHPLSPIQRPPKRHQEEVASVSSSS

351 KQKPVKMARRPTNLSPTALHKLITGGPSLESKSLTSEESAKLGWNANATE

401 ISDDTMSKLRGIVSEEVGTILRALERGRLKIVPSRSERMEMSPTGSRSSS

451 RNDKAPAVKDVEPTVPDTVPTEIISLSLRKDVQKAEVGIEPTIPDTASET

501 TSVST#

Figure 6.1. Sequence characterization of MCNA protein mcnA encodes for a 506 amino acid protein with a molecular weight of 56.6kDa. The protein contains a globular domain (2-270 aa), a coiled-coil domain (120-150 aa), four APC degradation box motif with RXXL sequence (88-93 aa, 167-172 aa, 187-192 and 425-430 aa, in red), substrate recognition site that interacts with cyclin thereby increasing phosphorylation by CDK/cyclin (RLLEL sequence, 51-55 aa) and four potential CDK1 phosphorylable consensus “SP” sites (underlined). 142

Figure 6.2. Sub-cellular localization of MCNA (A) A single fluorescent focus of MCNA-GFP in individual conidia was observed. (B) Western blot analysis was performed to analyze MCNA-GFP protein. MCNA-GFP is predicted to be 56.6 kDa but appears to migrate as several specific molecular species. (C) When GFP tagged cells are grown, fixed and stained with DAPI, a single MCNA body (arrowhead) is observed to appear within the nuclei in about 90% of the cells. (D) When the ER is visualized using a red fluorescent tagged ER marker Erg4, MCNA- GFP localizes near the nuclear periphery.

143

Figure 6.3. MCNA-GFP localizes in the vicinity of the nucleolus (A) When MCNA-GFP tagged cells are fixed and stained for ANA1, one can visualize the MCNA dot near the nucleolus. The MCNA-GFP body (green) locates next to the nucleolus (red). DNA is stained with DAPI (blue). (B) Three different nucleolar proteins, Fibrillarin, Bop1 and CgrA, endogenously tagged with red fluorescence protein, when visualized along with MCNA-GFP, clearly show MCNA to localize to a sub-domain of the nucleolus.

144

Figure 6.4. MCNA through the cell cycle (A) A cell containing 4 MCNA bodies was visualized through a single cell cycle and seen to double in number (The two bodies to the extreme right are separated from the rest by a septa and have exited the cell cycle). In the process of doubling, they undergo a unique pattern of segregation, highlighted by the appearance of multiple foci. (B) MCNA-GFP (green) in relation to chromatin (red). The beginning of M phase is marked by the condensation of chromosomes (panel a) followed by DNA segregation into two daughter nuclei (panel b). During condensation, a single MCNA body remains associated with the nucleus but as the DNA separates (panel a), the MCNA body is excluded from the nucleus and appears in the cytoplasm (panel b, arrowhead). About 10 mins after completion of mitosis MCNA protein begins to appear in the daughter nuclei (panel c). The single MCNA body at this time also divides into variable numbers of bodies, usually 2-4 (panel d), which linger in the cytoplasm for a given amount of time (panel e). After ~35 mins of its first appearance in the daughter nuclei MCNA bodies completely disappear from the cytoplasm (panel f). 145

B

MTATLERSLSRTSSVSMPLSPMISFRHEATTPISESSLSHIHERLSAIES RLLELRSTALTKDGYVDRRNREDEHIRREFEAHRSISNRIDLNVVALRAD VGQLKSGILQLKSSLGQAGNETVFLRSDVDRLSKNVDQIQVDLEHMQTDV CGCRVEISKLQSAISQLRTELITLQHETSRHLNSFLERFSLIEARMKHSE RVRFNSLAHTTHAPITPVPIVEEDGSLRWPDYFPRTVWRFWCLKKRSRHN RLAQLAEFYQLGGYEYWGRMHQTDGMFSDSDSSDSSDCPSNLTRAEAVRM FPEAAHQALAATLGLVYYKIRNEVGDHPLSPIQRPPKRHQEEVASVSSSS KQKPVKMARRPTNLSPTALHKLITGGPSLESKSLTSEESAKLGWNANATE ISDDTMSKLRGIVSEEVGTILRALERGRLKIVPSRSERMEMSPTGSRSSS RNDKAPAVKDVEPTVPDTVPTEIISLSLRKDVQKAEVGIEPTIPDTASET TSVST

Figure 6.5. MCNA is a phosphoprotein (A) Endogenously S-tagged MCNA protein after in vitro phosphatase treatment. Lane A shows protein with no protein phosphatase, Lane B is with added protein phosphatase and Lane C is control lane with additional of phosphatase and phosphatase inhibitor. Note the collapse in lane B of MCNA protein bands into a single band which proves that MCNA is modified by phosphorylation. (B) Mass spectrometric analysis of MCNA. Sequence in red shows sequence coverage while amino acids (serine 330, 365 and 442) shown in blue were identified as phosphorylation sites.

146

Figure 6.6 Characterization of mutant MCNA-3A (A) Mutant MCNA-3A-GFP with three phosphorylation sites mutated to alanine was analyzed by western blot using GFP antibody. Note the appearance of two molecular weight bands suggesting existence of additional phosphorylation sites in the protein not targeted by the mutagenesis. (B) MCNA-3A-GFP visualized in asynchronously grown cells fixed and DNA stained by DAPI. Note the unequal amounts of MCNA-3A-GFP in individual nuclei (right panel) compared to wild type MCNA-GFP cells (left panel). (C)Western blot analysis of protein made from MCNA-GFP strain and mutant MCNA- 3A-GFP strain using GFP antibody shows up-regulation of MCNA-3A protein in comparison to MCNA.

147

CHAPTER 7

DURING MITOSIS THE NUCLEOLUS OF ASPERGILLUS NIDULANS UNDERGOES STEPWISE DISASSEMBLY IN THE CYTOPLASM AND IS REASSEMBLED IN G1 NUCLEI

7.1. Introduction Cell division requires the duplication of all critical cell components, their distribution into different domains within the cell, and the separation of these domains into two independent daughter cells. These events are best characterized for DNA, the cells genetic material, which is duplicated during the S phase of the cell cycle, segregated into chromatids during mitosis, followed by formation of two daughter cells by cytokinesis. The membrane bound organelles of the eukaryotic cells can be expected to follow a similar route of segregation, the specific details of which would be determined by their functional requirements and their structural organization in the cell. The highly regulated manner of chromosomal segregation into two daughter cells require that they be attached to the mitotic spindle which emanates from the spindle pole bodies located at opposite ends of the cell. Before segregation is attempted, checkpoints ensure the completion of two events, one that the chromosomes are duplicated and two that they are attached in a bipolar fashion to the spindle poles. It is believed that the segregation of organelles like mitochondria and Golgi stacks, involves their sorting to distinct domains of the cytoplasm where their functions are most needed during cell division before being separated into the two daughter cells (Nebenfuhr, Frohlick et al. 2000). Their dependence on the spindle for equal separation however remains undetermined. On the other hand, the bipolar nature of the mitotic spindle guarantees the equal segregation of the spindle bodies themselves (Jaspersen and Winey 2004).The mitotic segregation of the membrane

148 less organelle, the nucleolus, however has been a less clear subject. Not much is known about the mechanism of nucleolar segregation or about its regulation during mitosis (Boisvert, van Koningsbruggen et al. 2007). The nucleolus as mentioned in section 1.5 is the site of rRNA genes and provides a structure which encompasses all the steps associated with ribosomal biogenesis (Perry and Errera 1961; Perry 1962; Brown and Gurdon 1964; Tschochner and Hurt 2003). The chromosomal rRNA genes form the nucleolar organizer which is alternatively called the nucleolar organizing region (NOR) (Heitz 1931; Heitz 1933). Using genetic, proteomic and bioinformatics approaches, more than 700 proteins in human and 200 proteins in plants have recently been shown to be associated with the nucleolus (Andersen, Lyon et al. 2002; Scherl, Coute et al. 2002; Leung, Andersen et al. 2003; Andersen, Lam et al. 2005; Coute, Burgess et al. 2006; Staub, Mackowiak et al. 2006). Other characterized nucleolar proteins have been shown to localize to three morphologically distinct regions of the nucleolus depending on their function during ribosome formation (Boisvert, van Koningsbruggen et al. 2007). The transcriptional proteins localize to the center (fibrillar center) of the nucleolus, pre-rRNA processing proteins surround the fibrillar center (dense fibrillar component) and ribosomal subunit assembly occurs at the periphery of the nucleolus (granular component) (Shaw and Doonan 2005). The segregation of the nucleolus during mitosis has been studied both in higher and lower eukaryotic cells and the location of nucleolar components including the NOR during the different phases of mitosis appear to be highly different between the two. In both Saccharomyces cerevisiae and in Saccharomyces pombe, the nucleolus remains intact during mitosis, ultimately being separated with the NOR along the mitotic spindle. In mitotic S. pombe cells and S. cerevisiae cells, the nucleolus trails the bulk of the DNA (Granot and Snyder 1991; Fuchs and Loidl 2004). In contrast, the nucleolus from higher eukaryotes undergoes disassembly during mitosis and various nucleolar proteins leave the structure in an orderly fashion while the NOR segregates with chromosomes. The mitotic disassembly of the nucleolus therefore requires mechanisms by which the entire nucleolus is reassembled during exit from mitosis and entry into a new cell cycle. These fundamental differences regarding nucleolar segregation between lower and higher

149 eukaryotes most likely can be attributed to the type of mitosis they undergo. The unicellular yeasts have a closed mitosis with an intact nuclear envelope while higher multicellular organisms undergo an open mitosis which involves nuclear envelope breakdown (Rabut, Lenart et al. 2004; Desouza and Osmani 2007). In S. cerevisiae, the stretch of rDNA gene arrays that constitutes the NOR are anywhere from 100 to 200 units of a 9.1kb repeat (Machin, Torres-Rosell et al. 2005). Studies show that late anaphase compaction of the rDNA is necessary for its proper segregation (Machin, Torres-Rosell et al. 2005). The compaction of the rDNA is in addition to sister chromatid compaction that occurs during normal DNA condensation at prophase. This is to ensure full segregation of the NORs independent of the spindles as anaphase spindles are thought to be insufficiently long for the separation of the chromosome containing the rDNA arrays (Machin, Torres-Rosell et al. 2005). The rDNA hyper compaction is regulated by the Cdc14 protein phosphatase, a key regulator of yeast mitotic exit (Garcia and Pillus 1999; Shou, Seol et al. 1999; Visintin, Hwang et al. 1999; Machin, Torres-Rosell et al. 2005) and is known to require the condensin protein complex (Freeman, Aragon-Alcaide et al. 2000). In spite of the change in chromosomal rDNA organization, no report of mitotic nucleolar disassembly has been reported in the yeasts. During open mitosis, the dynamic disassembly of the nucleolus is initiated by the transcriptional inactivation of the rRNA genes (Boisvert, van Koningsbruggen et al. 2007). The binding of the RNA pol I transcription factor SL1 (selectivity factor 1) and upstream binding transcription factor (UBF) at the rRNA gene promoter is required for the transcription of rRNA genes (Comai, Zomerdijk et al. 1994; Lin, Navarro et al. 2006) and has been shown in vitro to be regulated by CDK1/cyclin B directed phosphorylation of SL1 to prevent its association with UBF leading to its dissociation from the NOR (Heix, Vente et al. 1998; Kuhn, Vente et al. 1998). Although some components of the rDNA transcriptional machinery are removed from the NOR others, including RNA Pol I, UBF and Topoisomerase I, remain at the NOR throughout mitosis (Christensen, Larsen et al. 2002; Leung, Gerlich et al. 2004). On the other hand, all proteins involved with rRNA processing and ribosomal maturation are thought to be completely disassembled from the

150 NOR during initiation of mitosis (Leung, Gerlich et al. 2004). These disassembled nucleolar proteins are either located on the chromosome periphery from late prophase to early telophase and include fibrillarin (Yasuda and Maul 1990; Gautier, Fomproix et al. 1994; Scheer and Weisenberger 1994), nucleolin (Scheer and Weisenberger 1994) and B23 (Ochs, Lischwe et al. 1983) or are uniformly distributed in the cytoplasm (Jimenez- Garcia, Segura-Valdez et al. 1994) and include ribosomal protein S6 (Jimenez-Garcia, Segura-Valdez et al. 1994) and phosphoprotein p130 (Pai, Chen et al. 1995). Location of the rRNA-processing proteins to mitotic chromatin may help ensure these proteins are segregated evenly between daughter nuclei. Nucleolar reformation starts in early telophase when partially processed rRNA transcripts, together with specific nucleolar proteins, begin to form structures termed prenucleolar bodies (PNBs) in the new daughter nuclei (Jimenez-Garcia, Segura-Valdez et al. 1994; Dundr, Misteli et al. 2000; Savino, Gebrane-Younes et al. 2001). When RNA Pol I transcription is reinitiated at the chromosomal NORs in late telophase the PNBs are targeted to the NORs where they coalesce reforming daughter nucleoli (Scheer, Thiry et al. 1993; Jimenez-Garcia, Segura-Valdez et al. 1994; Scheer and Weisenberger 1994). Studies show that several non-ribosomal proteins also accumulate in large cytoplasmic particles termed nucleolus-derived foci (NDF) during anaphase and telophase (Dundr, Leno et al. 1996). These foci are thought to be an intermediate location of nucleolar components between the chromosomal periphery and the PNBs (Dundr, Meier et al. 1997). To monitor nucleolar reassembly in live cells, dynamics of fluorescently tagged nucleolar proteins have been visualized. These data demonstrate that proteins belonging to the granular component re-localize later than those of the dense fibrillarin component, while proteins of the dense fibrillar component relocalized later than those in the fibrillar component (Dundr, Misteli et al. 2000; Savino, Gebrane-Younes et al. 2001; Leung, Gerlich et al. 2004). This stepwise reassembly of proteins into growing nucleoli is thought to reflect the order in which their functions are required for restarting ribosomal biogenesis. This order is also reflected in the reformation of the nucleolar substructure (Leung, Gerlich et al. 2004).

151 Among regulators of nucleolar disassembly and reassembly during mitosis, type 1 protein phosphatase (PP1) is considered to be a potential candidate. PP1 catalyses the removal of phosphate residues from serine and threonine and are among the most highly conserved proteins in eukaryotes. Mutations in the PP1 gene of S. cerevisiae, glc7, causes cells to arrest before anaphase with replicated DNA and duplicated spindle poles due to activation of the spindle assembly checkpoint, resulting in reduced cell viability (Andrews and Stark 2000). Similarly, the bimG11 mutant of A. nidulans causes an anaphase defect, with cells containing duplicated spindle poles, short aberrant spindles and unseparated DNA (Doonan and Morris 1989). Additionally, in the bimG11 mutant the nucleolus persists till late mitosis with elevated levels of phosphorylated nucleolar proteins as measured by MPM2 phospho specific antibody staining (Fox, Hickey et al. 2002). Data collected after the loss of BIMG activity thus suggests that the phosphatase may play a direct role in nucleolar dynamics, potentially being required to remove phosphate groups from phosphorylated nucleolar proteins during exit from mitosis. Moreover, BIMG-GFP locates to several specific sub cellular locations, including potentially the nucleolus late into mitosis (Fox, Hickey et al. 2002). These data are suggestive of a functional role for BIMG mediated dephosphorylation during segregation of the nucleolus during A. nidulans mitosis. The mitotic segregation of the nucleolus in Aspergillus nidulans is of particular interest from an evolutionary perspective as this mitotic system is considered neither to be open nor closed but in between. A. nidulans undergoes a partial open mitosis (Desouza and Osmani 2007) with no nuclear envelope breakdown but involving the partial disassembly of the nuclear pore complex. The mitotic opening of the NPC allows cytoplasmic proteins into the nucleus and access of the nucleolus directly to the cytoplasm. This chapter investigates the mode of nucleolar segregation in A. nidulans partially open mitosis. The findings most surprisingly define how the nucleolus is first expelled into the cytoplasm during mitosis and how it is subsequently disassembled and reassembled into daughter nuclei during exit from mitosis in a step-wise manner. I will also present data regarding regulation of this process.

152 7.2. Results 7.2.1. Mitosis in A. nidulans involves a double pinch of the nuclear envelope resulting in the formation of a transient “nuclear remnant”. Live cell imaging of certain nuclear pore complex proteins (Nups) during mitosis revealed that nuclear partitioning of the nuclear envelope (NE) to form daughter nuclei may involve a double pinch in A. nidulans (Osmani, Davies et al. 2006). This phenomenon is best observed with the Nup An-Gle1-GFP, see supplementary movie 7 in (Osmani, Davies et al. 2006). An-Gle1-GFP remains at the NE throughout mitosis and clearly locates around three structures during late mitosis (Figure 7.1A, panel b). The central structure is transiently present for a short period of time and ultimately disappears while the two structures on either side go on to become daughter nuclei (Figure 7.1A, panel c). This indicates that two nuclei are generated from one parental nucleus via a double pinch mechanism rather than a single pinch mechanism previously defined for the closed mitoses of Saccharomyces cerevisiae and Schizosaccharomyces pombe (Rose, Misra et al. 1989; Hurt, Mutvei et al. 1992; Copeland and Snyder 1993; Koning, Lum et al. 1993; Yeh, Skibbens et al. 1995). The double pinch mechanism results in the formation of a transient “nuclear remnant” structure. The double pinch phenomenon was also observed while visualizing certain Nups which remain at the NE throughout mitosis such as Nup96 (Ukil, De-Souza et al. In preparation). However, not all Nups that remain during mitosis are located to the central structure, for example, membrane Nup An- Pom152 is shown to locate to the daughter nuclei structures alone (Ukil, De-Souza et al. In preparation). More than half of the known Nups of A. nidulans are dispersed from the NPC during mitosis but return to the NPC during mitotic exit (Osmani, Davies et al. 2006). A preferential return of the dispersed Nups to the structures destined to become daughter nuclei was observed (Ukil, De-Souza et al. In preparation). Cells expressing Nup96- mRFP (non-dispersed Nup that remains at NPC through mitosis) and GFP-Nup98 (dispersed Nup during mitosis) were fixed and visualized for mitotic figures containing the nuclear remnant structure. In contrast to Nup96-mRFP, which surrounded the nuclear remnant, GFP-Nup98 localized to the daughter nuclei alone (Ukil, De-Souza et al. In

153 preparation). This suggests that mitotically dispersed Nups have a preference for the NE located around the daughter nuclei during nuclear pore complex reassembly post mitosis over the NE surrounding the nuclear remnant. Preferential re-assembly of the Nups most probably is a reflection of the properties of the newly forming daughter nuclei. Because the outer nuclear membrane is contiguous with the ER, fluorescent markers of the ER can also be used to visualize NE behavior (Zweytick, Hrastnik et al. 2000). The visualization of an ER marker protein An-Erg4 also therefore allows the observation of double pinching of the nucleus during mitosis (Figure 7.1B). The An- Erg4-GFP signal revealed the distribution of the ER within the cytoplasm and around nuclei in the same pattern as previously reported for another ER marker in A. nidulans (Fernandez-Abalos, Fox et al. 1998). Importantly, during mitosis the An-Erg4-GFP signal around the NE displayed very similar behavior to An-Gle1 and clearly revealed the existence of the three “nuclear” structures (Figure 7.1B, arrowhead) (Ukil et al. unpublished data). The nuclear remnant was confirmed to be a very transient structure (~2 minutes) and on occasion appeared to be reabsorbed into one of the daughter nuclei. In the majority of cases the remnant disappeared, presumably by reincorporation into the ER.

7.2.2. The nuclear remnant is devoid of DNA To see if the nuclear remnant contained any nuclear DNA we visualized both the ER (Erg4-GFP) and nuclear DNA (histone H1-mRFP) during progression through mitosis (Figure 7.1C). In late G2, nuclear DNA was not condensed and was surrounded by the NE (Figure 7.1C, panel a). As cells entered mitosis nuclear DNA became condensed and sister chromatids began to separate at anaphase and continued to separate during telophase (Figure 7.1C, panel b). At the same time the nuclear remnant was formed in between the separated condensed nuclear DNA (Figure 7.1C, panel b). Noticeably the remnant, although surrounded by NE, did not contain nuclear DNA. Upon completion of mitosis, and entry into G1, the nuclear DNA decondensed and the nuclear remnant was no longer visible as cells reassumed their interphase configuration, but with double the number of nuclei ((Figure 7.1C, panel c).

154 7.2.3. The nucleolus of A. nidulans segregates during mitosis via a mechanism that generates three “nuclear” structures The nucleolus does not break down during the closed mitoses of S. cerevisiae and S. pombe but is segregated along with DNA on the mitotic spindle (Granot and Snyder 1991; Fuchs and Loidl 2004). This pattern of nucleolar segregation is not seen in A. nidulans. Instead, the parental nucleolus of A. nidulans undergoes disassembly at more or less the same time that new nucleoli are being reformed in daughter nuclei. This pattern of segregation can be seen for An-fibrillarin-GFP (Fib-GFP) during live cell imaging and is shown as panels in figure 7.2. During interphase, Fib-GFP locates almost exclusively to the nucleolus although a very minor amount can just be detected in the nucleoplasm, near the nucleolus (Figure 7.2A, panel a). During progression through mitosis the nucleolar signal of Fib-GFP diminishes with time whilst two new foci of Fib-GFP re- appear either side of the parental nucleolus (Figure 7.2A, panel c and d). The two new foci of Fib-GFP are presumably within the two newly generated daughter nuclei and constitute reforming daughter nucleoli (Figure 7.2A, panel e). To determine if the behavior of Fib-GFP during mitosis was unique to this nucleolar protein, and to define the relationship between nucleolar segregation and DNA segregation, four strains were generated. Each of these strains express histone H1-mRFP for visualization of DNA along with endogenously tagged GFP versions of nucleolar proteins An-Fib, An-CgrA, An-Bop1 or An-Nrap. Live cell imaging of each strain through mitosis yielded representative data shown in figure 7.2B and 7.3. For Fib-GFP with H1-mRFP individual time series are depicted along with merged series (Figure 7.2B). For the other three strains the merged series alone are shown (Figure 7.3). During interphase, the nucleolus typically resides in a sub-region of the nucleus (Figure 7.2B, panel a), often offset from the nuclear center, abutting a region of the NE. DNA is largely excluded from the nucleolar region (panel b). As DNA is segregated during anaphase and telophase Fib-GFP does not segregate with the DNA (panel c). Instead the Fib-GFP mass is left behind, in between the separated DNA (arrowhead, panel d). After DNA is completely segregated new nucleoli begin to appear within each segregated mass of DNA (panel e). The appearance of the expanding daughter nucleoli corresponds with a

155 proportional decrease in the parental Fib-GFP signal. With time the entire parental Fib- GFP signal is lost and all Fib-GFP becomes located within daughter nuclei. These G1 nuclei thus appear as smaller version of the starting G2 nuclei (panel f). During this process, as the parental nucleolus disappeared and the new nucleoli appeared, we could detect an increase in the cytoplasmic level of Fib-GFP. However, when the new nucleoli were completely reformed the cytoplasmic signal had completely diminished. The pattern of segregation of CgrA-GFP, Bop1-GFP and Nrap-GFP in relationship to segregation of DNA was identical to that of Fib-GFP (Figure 7.3). This indicates that the entire nucleolus is segregated in A. nidulans via disassembly and reassembly each nuclear division cycle. Several details of this process are worth pointing out. As mentioned above, the nucleolus is often offset from the nuclear DNA and is thus located to one side of the nucleus in late G2 prior to entry into mitosis. However, as mitosis proceeds the parental nucleolus always becomes positioned between the separating daughter nuclei. This location can be central, as can be seen for CgrA (figure 7.3, panel c), or can be more towards one daughter nucleus, as seen for Bop1-GFP (figure 7.3B, panel c). Another point of mechanistic interest is the fact that the cytoplasmic level of nucleolar proteins is exceptionally low during interphase but becomes slightly elevated during mitosis (Figure 7.2B, panel b, c, d and e). No obvious tracks, or streaming, of nucleolar proteins between the parental and daughter nuclei are observed. The cytoplasmic levels then decrease as daughter nucleoli finish reforming. These data suggests that the mechanism of redistribution of the parental nucleolar proteins to the daughter nuclei involves general release of nucleolar proteins from the parent structure to the cytoplasm followed by nuclear import of these released proteins into newly formed daughter nuclei during G1. The imported nucleolar proteins then reform new nucleoli.

7.2.4. The transient nuclear remnant formed by double NE pinch contains the nucleolus but not the nucleolar organizing region (NOR) We considered the possibility that the transient parental nucleolus formed during mitotic progression might be inside the nuclear remnant formed by the double pinching

156 of the NE. We visualized Erg4-GFP (ER) and Fib-chRED (nucleolus) to investigate this possibility. Live cell imaging revealed that the nuclear remnant formed by the double NE pinching contained the parental nucleolus (Figure 7.4A, arrowhead). As the parental nucleolus diminished the ER membrane surrounding it was reabsorbed and the membrane bound nucleolus disappeared. Thus the parental nucleolus resides in the nuclear remnant which transiently exists between daughter nuclei during exit from mitosis. The nucleolus is known to be formed around the NOR which consists of multiply repeats of the ribosomal genes (McClintock 1934; Batistoni, Andronico et al. 1978; Cheutin, O'Donohue et al. 2002). Because the parental nucleolus is separated from DNA during A. nidulans mitosis (Figure 7.2B) it is possible that the nucleolus can perhaps exist without the NOR, at least transiently. Alternatively the NOR might not segregate with the main mass of nuclear DNA but could perhaps segregate late during mitosis, as occurs in yeast systems. In mammalian cells topoisomerase 1 (Topo I) remains associated with the NOR throughout the cell cycle (Leung, Gerlich et al. 2004) and importantly some Topo I is maintained at the NOR during mitosis, even after the nucleolus has disassembled. We therefore endogenously tagged the Topo I gene of A. nidulans with GFP to act as a marker to follow the NOR during mitosis. We then defined the relationship between Topo I and mitotic chromatin using live cell imaging of double Topo I-GFP (NOR) and H1-mRFP (DNA) tagged cells (Figure 7.4B). During interphase and late G2 Topo I-GFP resides predominantly in the nucleus with some co-localizing in the vicinity of the nucleolus (Figure 7.4B, panel a). Upon entry into mitosis, before chromosome condensation, the majority of Topo I-GFP is released from nuclei but a small fraction remains within nuclei in an area offset from the main chromatin mass (panel b). As chromosome condensation proceeds Topo I-GFP becomes located to a single dot which likely represents the un-segregated NORs (panel b arrowed). In support of this idea, when anaphase fires the single Topo I-GFP dot is split in two and one focus is segregated to each daughter nucleus along with the segregating DNA (panel c). We did not observe any Topo I-GFP with lagging chromosomes during anaphase, as would be expected if the NOR was segregated late during mitosis, as occurs in yeast. As mitotic exit proceeds the nuclear levels of Topo I-GFP increases (panel d) and as cells pass into G1 the pre-mitotic

157 configuration of Topo I-GFP, in relationship with H1-mRFP within nuclei, is reestablished (panel e). As Topo I-GFP segregates with mitotic chromatin, and the nucleolus remains as a single entity after DNA segregation has occurred, this indicates the NORs are removed from the nucleolus at some time during progression through mitosis. To directly test this we followed the distribution of Topo I-GFP (NOR) in relationship with the Fib-chRED (nucleolus) during mitosis. In G2, when both Topo I-GFP and Fib-chRED are predominantly nuclear, there is some overlap between both proteins indicating Topo I- GFP is present both in the nucleus and the nucleolus (Figure 7.4C, panel a). As described above, upon entry into mitosis the majority of Topo I-GFP is released from nuclei, and at this point there is more obvious co-localization of Topo I-GFP with Fib-chRED inside nucleoli (Figure 7.4C, panel b). However, as the Topo I-GFP signal continues to decrease to become concentrated to a single dot, the dot resides towards the edge of the nucleolus (Figure 7.4 B, panel b). Most importantly, during anaphase, when the single dot of Topo I-GFP is split into two and segregates with DNA, (Figure 7.4B, panel c) the nucleolar Fib-chRED remains between the two separated Topo I-GFP foci (Figure 7.4C, panel c). As mitosis proceeds the nucleolus undergoes disassembly (Figure 7.4C, panel d). Upon entry into G1 the dispersed Fib-chRED starts to co-localize with the Topo I-GFP foci within the daughter nuclei and both nuclear signals continue to increase until their interphase configuration in daughter nuclei is reestablished ((Figure 7.4C, panel e). The fact that the new daughter nucleoli reassemble around the Topo I foci again indicates that the mitotic dots of Topo I correspond to the NORs.

7.2.5. Daughter nuclei re-establish transport before reassembling nucleolar proteins If nucleolar proteins are released from the parental nucleolus and re-imported to daughter nuclei, the appearance of nucleoli in G1 nuclei should occur after nuclear transport has been reestablished in G1. To investigate this we followed nuclear transport using an NLS-DsRed marker and formation of nucleoli with Fib-GFP (Figure 7.5). We consistently detect nuclear accumulation of NLS-DsRed slightly before (Figure 7.5, compare panel c and d), or at the same time as, nuclear accumulation of Fib-GFP (Figure

158 7.5, compare panel d and h). This indicts that nuclear transport is first established and then new daughter nucleoli are formed in G1. When we repeated this experiment using the nucleolar protein Bop1, we again consistently detected nuclear accumulation of NLS- DsRed before Bop1-GFP. However in this case, NLS-DsRed first appeared in the nucleus at least 4-5 minute earlier than Bop1. This suggests that the reassembly of Fib-GFP occurs earlier than Bop1-GFP.

7.2.6. Fibrillarin reassembles in daughter nucleoli earlier than Bop1 If nucleolar proteins are released from the parental nucleolus in a step-wise fashion and reassembled into daughter nuclei in a step-wise fashion, the dispersal of Fib from the parental nucleolus and the appearance of Fib in the daughter nuclei should occur earlier than Bop1. To test this we followed the nucleolar segregation of Fib-GFP along with Bop1-chRED. Fib-GFP was observed to be more in the center of the nucleolus as compared to Bop1-chRED which was to the periphery (Figure 7.6, Merge panel a). This localization pattern conforms well to where these proteins reside within the mammalian nucleolus. Fib is supposed to be an early processing protein localized to the dense fibrillar center while Bop1 is a late processing protein localized to the granular center. As mitosis proceeds, Fib-GFP can be seen to be the first to disassemble from the parental nucleolus resulting in an increase in the cytoplasmic Fib-GFP signal (Figure 7.6, panel b) while Bop1-chRED remains intact. Fib-GFP continues to disassemble and reassemble into the daughter nucleoli (a red bar next to the Fib-GFP series marks the disassembly- reassembly of Fib-GFP) with no change seen in the pattern of the Bop1-chRED signal. It is only after Fib-GFP almost completes its reassembly into the two daughters that Bop1- chRED begins to disassemble from the parental nucleolus (panel c). Bop1-chRED then completely disassembles from the parental nucleolus before being reassembled into the two daughter nucleoli (a green bar next to the Bop1-chRED series marks its disassembly- reassembly). Clearly, the nucleolar disassembly of Fib from the parental nucleolus precedes Bop1 and so does the reassembly of Fib to the daughter nucleoli precede Bop1. Thus, the nucleolus in A. nidulans undergoes a step-wise disassembly and a step-wise reassembly of its constituent proteins during mitosis and into G1.

159 Note the distinctive stretch in the nucleolar structure in panel b for both Fib-GFP and Bop1-GFP before it suddenly changes into a spherical shape. The stretch most probably reflects the removal of the NOR from the parental nucleolus along the mitotic spindle followed by the double-pinching of the membrane which limits the nucleolar proteins into the spherical nucleolar remnant structure. This structural change of the nucleolus was consistently observed in other nucleolar protein tagged strains as well.

7.2.7. Nucleolar disassembly-reassembly is under the control of the spindle assembly checkpoint (SAC) Treatment of cells with microtubule poisons does not prevent entry into mitosis but does prevent progression past metaphase due to activation of the SAC. We wished to determine if segregation of the nucleolus was under the control of the SAC. As defined above, the segregation of the nucleolus involves disassembly of the nucleolus in the cytoplasm followed by its reassembly in daughter nuclei. We therefore asked if disassembly and/or reassembly of the Fib-GFP and Bop1-GFP were prevented upon activation of the SAC. To monitor mitosis, and activation of the SAC, we employed strains expressing NLS-DsRed which provides a good marker for both the start (prophase) and the end (G1) of mitosis. Release of NLS-DsRed from nuclei marks entry into prophase whereas re-import of NLS-DsRed provides a precise marker for early G1 as mitosis is completed. We have found that nuclear re-import of NLS-DsRed during exit from mitosis is under control of the SAC (De Souza and Osmani, Unpublished). Thus continued dispersal of NLS-DsRed from nuclei in the presence of benomyl (microtubule poison) indicates the SAC is engaged. On the other hand, return of NLS-DsRed to nuclei in the presence of benomyl indicates the SAC to no longer be engaged. In a wild type strain, mitosis is completed in ~5 min during which NLS-DsRed is first dispersed and then imported to daughter nuclei. In addition, the nucleolus is disassembled and reassembled equally in daughter nuclei for both Fib-GFP and Bop1- GFP (Figure 7.7A and Figure 7.8A, respectively). However, when cells enter mitosis in the presence of benomyl, NLS-DsRed is dispersed and remains dispersed for an extended period of time due to activation of the SAC (Figure 7.7B and Figure 7.8B, respectively).

160 During this extended mitotic arrest we found that the nucleolar proteins behave slightly different based on the protein being followed. Fib-GFP undergoes a slight dispersal from the nucleolar structure causing a slight decrease in its signal (Figure 7.7B) but it does not undergo complete disassembly. Bop1-GFP on the other hand remains intact and does not undergo any dispersal (Figure 7.8B). This indicates that the disassembly of the nucleolus is controlled by the SAC. The SAC can be subverted by deletion of key components of the system, amongst which is Mad2. In a Mad2 deleted strain (ΔMad2), cells can enter mitosis in the presence of benomyl but cannot engage the SAC. Such cells therefore proceed through mitosis into G1 without completing nuclear division. In a ΔMad2 strain, entry into mitosis correlates with release of NLS-DsRed from nuclei, as normal, after addition of benomyl (Figure 7.7C and Figure 7.8C). However, rather than remaining dispersed, the NLS-DsRed is re- imported to nuclei indicating the cells have exited mitosis and entered G1 even though no nuclear division occurs. This demonstrates the SAC is overcome when Mad2 is deleted, as previously shown in many other systems. Because disassembly of the nucleolus is controlled by the SAC we expected the nucleolus to undergo some disassembly and therefore to detect Fib-GFP and Bop1-GFP released from the nucleolus and appear in the cytoplasm. Indeed, we could detect some release of Fib-GFP (Figure 7.7C, asterix) and Bop1-GFP (Figure 7.8C, asterix) into the cytoplasm, although only to very low levels. We had also anticipated that the released nucleolar proteins would then be imported back to the nucleus and reassembled into the nucleolus. In fact we did see this phenomenon but with the following interesting observation. As the parental nucleolus was disassembled a new separate nucleolus was reassembled at the same time for Fib-GFP (Figure 7.7C, panel a and inset) and for Bop1- GFP (figure 7.8C, panel a and inset). The series of events are as follows: (1) Cells enter mitosis and NLS-DsRed escapes from nuclei as nuclear transport is compromised when NPCs undergo partial disassembly. (2) The nucleolus remains intact for a period of time (~1-2 mins for Fib-GFP and ~4-5 mins for Bop1-GFP) and then begins to disassemble causing the cytoplasmic signal to increase slightly. (3) As the nucleolus begins to disassemble, nuclear transport is reestablished and NLS-DsRed is transported back into

161 nuclei. (4) The dispersed nucleolar protein is now re-imported and a new spatially separated nucleolus is formed (Figure 7.7C and 7.8C, inset, arrowhead points to old nucleolus and arrow marks the new nucleolus). Because dispersal of the nucleolus is normally prevented by activation of the SAC, but nucleolar disassembly occurs when Mad2 is deleted when the SAC should be activated, this data confirms that the complete disassembly of the nucleolus is under control of the SAC system. The data also suggest that a cycle of disassembly of the parental nucleolus and reassembly of a spatially separated “daughter” nucleolus can occur in the absence of spindle function. Given the normal physical separation of DNA from the nucleolus during mitosis, as revealed above, we had expected that disassembly of the old nucleolus and reassembly of a spatially separated new nucleolus would require mitotic spindle function. However, the disassembly and spatially separated reassembly of the nucleolus observed in benomyl when Mad2 is deleted (Figure 7.7C, 7.8C) indicates that this cycle can occur without the function of the spindle apparatus.

7.2.8. Non-degradable cyclin B prevents Bop1 disassembly Other levels of regulation that may play a role in nucleolar disassembly- reassembly may include phosphorylation driven by CDKs and the reversal of these phosphorylations by specific phosphatases. During the engagement of the SAC in the presence of Benomyl, the levels of CDK1- cyclin B are known to be maintained at an elevated level. Our data shows that at this period of time Bop1-GFP remains intact while Fib-GFP begins to undergo disassembly (Figure 7.9A). This suggests that the disassembly of Bop1 might be additionally under the control of CDK1 mediated phosphorylations. In order to test this possibility, Bop1-GFP disassembly was observed under in the presence of non-degradable cyclin B expression. Non-degradable cyclin B does not undergo degradation at mitotic exit thereby preventing CDK1-cyclin B inactivation. CDK1-cyclin B remains active through out and causes cells to arrest in late telophase. When Bop1-GFP was observed in telophase arrested cells after induction of non-degradable cyclin B expression, it was found to remain intact between the two masses of DNA as visualized by DAPI staining (Figure 7.9 B) and not undergo any

162 disassembly. The Bop1-GFP signal was sometimes located in the middle and sometimes offset to one of the DNA masses. However, no cytoplasmic signal for Bop1-GFP was observed. The quantified data is shown in table 7.1. When a similar experiment was done to observe the effect of non-degradable cyclin B on Fib-GFP, a different result was obtained. In the presence of active CDK1- cyclin B, cells had a telophase arrest with segregated masses of DNA (Figure 7.9C). However, Fib-GFP was seen to disassemble and a cytoplasmic signal observed. In some cases, reassembly of Fib-GFP to the daughter nuclei was also observed (Figure 7.9C and Table 7.1, marked as R). This data strongly suggests that CDK1-cyclin B activity plays a role in inhibiting Bop1-GFP disassembly but does not affect the disassembly of Fib-GFP.

7.2.9. The potential role of BIMG in nucleolar disassembly The potential role of CDK1 mediated phosphorylation in inhibiting nucleolar disassembly suggests the presence of a phosphatase that may counter remove the phosphorylations of CDK1 and cause nucleolar disassembly at mitotic exit. A potential candidate lies in the type 1 protein phosphatase BIMG. Localization studies of BIMG- GFP show it to be localized to various distinct sub-cellular localizations (Figure 7.10A). During interphase BIMG-GFP localizes to the spindle pole, slightly to the nucleoplasm, the germ tip and more importantly to the nucleolus (a crescent shape structure is observed within nuclei) (Figure 7.10A). During mitosis, BIMG is observed to go to the kinetochores (Figure 7.10B, panel a) followed by its localization to the nuclear remnant between the two separating spindle poles and co-localizing with Fib-chRED in the nuclear remnant (Figure 7.10B, panel b), before being reassembled to the daughter nuclei and reconstituting pre-mitotic sub-cellular localizations (Figure 7.10B, panel c). Note that the spindle pole localization of BIMG disappears specifically during prophase and metaphase and comes back during anaphase. Therefore, the distribution of BIMG in the cell and during mitosis suggests a potential role at the nucleolus and at the remnant. Previous studies with a mutant allele of BIMG, bimG11 has also shown cells to arrest in anaphase with duplicated spindle poles, unseparated DNA and an intact nucleolus with high levels of phosphoprotein staining as revealed by MPM2, a phospho specific

163 antibody. This suggests that the phosphatase may play a direct role in nucleolar dynamics, potentially being required to remove phosphate groups from phosphorylated nucleolar proteins during exit from mitosis.

7.2.10. The unique nucleolar segregation pattern of A. nidulans helps to ensure equal distribution of nucleolar proteins to daughter nuclei in a common cytoplasm. Aspergillus nidulans is a multi-nucleate organism containing multiple nuclei in a common cytoplasm, separated into cell compartments by the septa. Nuclei in a given cell compartment undergo nuclear division in a semi-synchronous manner meaning that nuclear division starts as a wave from the spore end of the cell to the germ tube end resulting in nuclear division in almost the same time. During mitosis when the NPC complex partially disassembles and the nuclear pores open, many soluble proteins and RNA equilibrate across the cytoplasm and nucleoplasm. Certain proteins remain associated with the DNA and the NE and segregate along the mitotic spindle. Proteins that leak into the cytoplasm and contain a nuclear import sequence (NLS) are re-imported at G1 when the NPC is reassembled and transport is turned back on. Similarly, proteins with a nuclear export sequence (NES) are exported out of the nucleus. We have used NLS-DsRed as a marker for nuclear transport. At prophase when the NPC partially opens, NLS-DsRed leaks out of the nucleus and is uniformly distributed through out the cell. When cells exit mitosis and reassemble the NPC, NLS- DsRed is localized back to the nucleus by active transport. However, since A. nidulans undergoes a semi-synchronous mitosis we observed that certain nuclei enter and exit mitosis slightly earlier than others (Figure 7.11, nucleus 1). These nuclei thus re- accumulated cytoplasmic NLS-DsRed early as well (daughter nuclei 1a and 1b). This resulted in a variation of NLS-DsRed amounts in individual nuclei (Compare daughter nuclei of 1 and 2). Nuclei resuming transport early have more NLS-DsRed in them while nuclei resuming transport slightly later had less NLS-DsRed in them due to a lack of the protein in the cytoplasm that can be imported. Therefore, a competition exists between the nuclei for cytoplasmic distributed NLS proteins that depend on active transport as their mode of re-accumulation. Surprisingly, when we observed the same set of nuclei for

164 the reassembly of nucleolar proteins we did not see any significant variation in the amounts of nucleolar proteins in the daughter nucleoli (Figure 7.11, Fib-GFP panel). This led us to believe that the segregation of the nucleolus during mitosis via a remnant may be of significance for equal distribution of the nucleolar proteins. Certain observations add value to this idea. 1) As mentioned above, nuclei entering and exiting mitosis at different time points still accumulate equal amounts of nucleolar protein. 2) The nucleolus is always in the middle of the two newly forming daughter nuclei. Even if the parental nucleolus is offset to begin with, it always re-localizes to the middle which may aid the daughter nuclei during reassembly by providing equidistant opportunity to import proteins from the remnant and reducing competition from neighboring nuclei. 3) The nucleolus undergoes a slow step-wise disassembly and does not distribute all of its constituent proteins into the cytoplasm in one go. The timing of disassembly is also closely tied with the resumption of active transport allowing quick localized reassembly of proteins from the remnant to its respective daughter nucleoli. In order to test the hypothesis that a nuclear remnant containing the nucleolus allows equal distribution of nucleolar proteins to daughter nuclei on either side of it, a mathematical approach was taken. Mathematical modeling of both proteins, one that disperses to the entire cytoplasm at mitosis and one that is retained in a remnant and slowly dispersed, was done using four parameters: the rate of nuclear import of proteins (values measured using NLS-DsRed data), the rate of nucleolar protein dispersion (values measured using Bop1-GFP data), diffusion constant and the time differential of mitotic entry and exit between nuclei undergoing semi-synchronous mitosis. All values were obtained from data collected during live cell imaging of nucleolar segregation while the diffusion constant value was taken from literature.

7.2.11. Mathematical modeling of cytoplasmic dispersed protein reassembly into daughter nuclei in a common cytoplasm The re-accumulation of a dispersed protein (like NLS-DsRed) from the cytoplasm into four daughter nuclei was modeled first and the output is shown as graphs in figure 7.12A. Graph A shows the total amount of dispersed protein in the cytoplasm after NPC

165 disassembly (green curve) of two parental nuclei. In graph B, one of the parental nuclei (we will call this N1) is modeled to exit mitosis and its daughter nuclei begin to re-import the dispersed protein from the cytoplasm which can be seen as two blue peaks. Note the dip in the green curve which represents the lowering of cytoplasmic concentrations of the protein. Daughter nuclei of N1 continue to reabsorb and this result in further lowering of the cytoplasmic levels of the protein (green curve in graph C and D) and a proportionate increase in the two blue peaks. After 40s of time, the second parental nucleus (N2) exits mitosis and is modeled to begin re-absorption. The second set of two blue peaks is representative of this (graph E). With time the dispersed protein amounts in the cytoplasm reach zero and re-absorption is complete. Note that the modeling of a dispersed protein gives the same result as live cell imaging of NLS-DsRed and accumulates more protein in the daughters of N1, the nucleus to exit mitosis first, compared to daughter nuclei of N2. Thus, the final amount of nuclear accumulation of a cytoplasmic dispersed protein is influenced by the timing at which import begins for individual nuclei.

7.2.12. Mathematical modeling of nuclear remnant protein reassembly We next modeled the nuclear accumulation of a protein that during mitosis is localized to a cytoplasmic nuclear remnant structure similar to nucleolar proteins and not generally dispersed into the cytoplasm. The data is represented in figure 7.12B as several graphs. In Graph A, the modeling begins with two parental nuclei (N1 and N2) leaving behind a high amount of protein localized to a remnant like structure in the cytoplasm, represented here as two red peaks. There is no cytoplasmic dispersion at this point of time. In graph B, N1 exits mitosis first and begins to form daughter nuclei and re- establish active transport. At the same time, a slow dispersion of the remnant protein into the cytoplasm begins. This can be seen as a very low increase in the cytoplasmic level of the protein (green curve). With time the daughter nuclei of N1 re-accumulate the remnant protein in them and this can be visualized as the two rising blue peaks on each side of the remnant red peak. The daughter nuclei continue to accumulate the remnant protein and the blue peaks continue to rise representative of increased protein accumulation with the

166 simultaneous decrease in the remnant red peak. After 40s, N2 is programmed to exit mitosis and its daughters to begin active transport. Concomitantly, the remnant protein of N2 begins to disperse in graph E and two blue peaks corresponding to daughter nuclei of N2 begin to appear. The remnant protein corresponding to N2 now continues to disperse and the daughter nuclei of N2 continue to re-accumulate. Again this can be seen as a decrease in the red peak and an increase in the neighboring blue peaks. Note that the two pairs of daughter nuclei maintain the time differential in terms of amount of remnant proteins that remains to be reassembled into their daughter nuclei. However, when the amount of remnant protein from each red peak reaches zero, the final amount of protein among the four daughter nuclei remain relatively equal. Therefore the modeling of a protein similar to nucleolar protein suggests that localized retention of protein (in a remnant) and its slow release between its target daughter nuclei coinciding with resumption of nuclear transport allows for more equal segregation of remnant proteins to daughter nuclei in spite of one nucleus exiting mitosis earlier than the other (N1 exits mitosis before N2). Thus the specific pattern of nucleolar segregation seen in A. nidulans most probably serves the purpose of equal nucleolar protein segregation to daughter nucleoli.

7.3. Discussion Mitosis, a critical and highly orchestrated event in the cell cycle, defines how cells divide and transmit genetic information from one cell generation to the next. Chromosomes are segregated in a highly regulated manner to ensure daughter nuclei are genetically identical and this is achieved by sister chromatids being attached to the mitotic spindle via kinetochore microtubules from the two spindle poles. Many years of work have led to a clear understanding of the many structural changes of the chromosomes during mitosis and the role of the mitotic spindle in chromosome segregation. For example, histone modifications play pivotal roles in conferring structural diversity to chromosomes by influencing the compactness of chromatin so that they can be easily segregated during mitosis with no physical chromosomal loss. Mitosis, however involves more than the segregation of chromosomes and spindle pole bodies alone. In

167 organisms like yeast and Aspergillus nidulans, the chromosomes are divided within the confines of the nuclear envelope. These organisms thus also ensure coordination between segregation of their genetic material with the division of the NE. Other sub-organelles, like the nucleolus and its constituents, also require coordinated mechanisms ensuring their equal division to the two daughter nuclei. The results in this chapter clearly establish a novel mechanism through which the nucleolus in A. nidulans segregates during mitosis and is shown to involve the expulsion of the nucleolus to the cytoplasm. Segregation of the nucleolus is also accompanied by major structural and functional changes to the NE and its distinct membrane domains including the nuclear pore complex. This section will discuss the new insights into the mechanism and various aspects involved in the mitotic segregation of the nucleolus and its interaction with the NE, explain possible regulatory mechanisms that ensures equal segregation and suggest reasons for A. nidulans undergoing such a novel and different type of mitotic nucleolar segregation.

7.3.1. The mitotic dynamics of the NE in A. nidulans The double layered nuclear envelope uniformly surrounds the chromosomes throughout the cell cycle. The NE composition however differs between interphase and mitotic nuclei. Beginning during mitosis, the NE is first modified by the onset of NPC disassembly resulting in structural changes that alter the permeation capability of the NE and allow the entry of soluble proteins and RNA molecules resulting in equilibration between the cytoplasm and nucleoplasm. Once anaphase is over the NE undergoes a double pinch resulting in the division of the NE into three structures distinctly defined by different forms of the NE. The two outer structures go on to become daughter nuclei and contain a form of NE functionally equivalent to the parental NE. The NE of the central structure is also similar to normal NE as it contains the ER marker Erg4 and other NPC proteins like Gle1, Nup96, and Nup120. However, there are certain differences that make the NE in the central structure differ from the NE present in the two outer structures. It contrast to normal NE, the central structure 1) is unstable and thus transient in existence, 2) is devoid of certain Nup proteins such as the integral membrane protein Pom152, 3) does not have

168 dispersed NPC proteins return to its NE which preferentially re-assemble to the outer structures instead and 4) is incapable of re-establishing transport of NLS containing proteins unlike the outer two structures. The process of generating two daughter nuclei in A. nidulans involves a double pinch of the NE unlike a single pinch of the NE in organisms like S. pombe and S. cerevisiae. Additionally, as a result of the double pinch a transient nuclear remnant structure is formed that is distinct from the two daughter nuclei. The formation of such different functional forms of NE structures from a single nucleus in a common cytoplasm is very fascinating and begs the question as to what may be the functional significance of the nuclear remnant and what may the nuclear remnant contain. Is it possible that the nuclear remnant is a functional equivalent of the midbody found in yeast and mammalian cells during cytokinesis that are thought to play a role as a dumping ground for unwanted proteins? Further studies of the contents of the nuclear remnant revealed it to contain nucleolar associated proteins.

7.3.2. The transient nuclear remnant defined by the modified NE contains the nucleolus The nucleolar segregation of A. nidulans when observed by visualizing tagged nucleolar proteins like Fibrillarin revealed a very unique pattern of segregation. The protein underwent disassembly at more or less the same time that new nucleoli were being reformed in daughter nuclei and this pattern is very different to the pattern of segregation observed in yeast, in which, the nucleolus segregates with DNA on the mitotic spindle. In order to directly test if the nucleolus in A. nidulans segregated with the DNA, Fibrillarin was visualized with DNA and discovered to behave in a completely unexpected manner. Fibrillarin does not segregate with DNA. Instead, as DNA segregates during anaphase and telophase, Fibrillarin is left behind in between the two separated DNA masses. It is only after DNA segregation is completed that new nucleoli, marked by Fibrillarin, begin to appear within each segregated mass of DNA. The appearance of the expanding daughter nucleoli corresponds with a proportional decrease in the amount of Fibrillarin located between the DNA masses. With time all Fibrillarin protein is

169 assimilated within the two daughter nuclei. This pattern is not specific to Fibrillarin alone and is reproducible for other nucleolar proteins located at different sub-nucleolar regions. During the process of nucleolar segregation, an increase in the cytoplasmic level of nucleolar proteins is detected. However, when the new nucleoli are reformed the cytoplasmic signal diminishes. If nuclear transport using a NLS-DsRed marker is followed, DsRed-NLS is seen to escape from the nucleus during mitotic initiation. This is due to the opening of the NPC. During G1, when the nuclear pore proteins are reassembled at the NPC, nuclear transport resumes and NLS-DsRed re-accumulates in the nucleus. The resumption of nuclear transport at G1 is probably required for the uptake of nucleolar proteins by active transport as the appearance of nucleolar proteins in the nucleus always occurs after nuclear transport is resumed. Data suggests that the mechanism of redistribution of nucleolar proteins to the daughter nuclei involves disassembly and general release of nucleolar proteins from the nucleolus to the cytoplasm followed by its reassembly via nuclear import into the two daughter nuclei then nucleoli. The appearance of the parental nucleolus and the mechanics of its disassembly and reassembly also suggested that the nucleolus may be inside the nuclear remnant earlier defined by the NE. By observing the NE and the nucleolar proteins at the same time, the idea of the nuclear remnant containing the nucleolus was confirmed.

7.3.3. Regulation of disassembly and reassembly of the nucleolus The pulling of the NOR from the nucleolus along the spindle is most probably a part of the nucleolar disassembly regulation. During mitosis there is a consistent change in the shape of the nucleolus. During chromosomal segregation the nucleolus appears as a long stretched rod shaped structure which quickly changes into a spherical shaped structure when DNA segregation is completed. This most probably reflects the pulling of the NOR from the nucleolus followed by the double pinch of the NE membrane resulting in the formation of the nuclear remnant. When the location of the NOR is visualized in combination with DNA, using Topo I-GFP and H1-mRFP, the NOR indeed leaves the nucleolus with DNA and does not localize in the remnant structure. The removal of the

170 NOR can thus result in a cascade of events beginning with the inactivation of RNA Pol I driven transcription of rRNA genes. The inactivation of RNA Pol I in other systems has been shown to initiate a structural change of the nucleolus and this may be the trigger for nucleolar disassembly in A. nidulans as well. Similar to higher eukaryotic open mitosis, certain components of the rRNA gene transcription machinery, like RNA Pol I and Topo I, remain at the NOR and are removed from the nucleolus along with the NOR. This most likely serves to promote nucleolar reassembly by acting as a seed for the reformation of the nucleolus. Physical removal of the NOR however cannot represent the entire regulatory system of nucleolar disassembly as the removal of the NOR does not result in an instant collapse of the entire nucleolus. Instead, proteins from the nucleolus undergo step-wise disassembly followed by a step- wise reassembly onto the daughter NORs. This ordered process suggests that additional regulatory steps are required for the disassembly of the nucleolus. Additional regulation of the nucleolus can exist in the form of CDK1 mediated phosphorylation counter balanced with phosphatase mediated dephosphorylation of key proteins. If this level of regulation exists then we would expect the nucleolar disassembly-reassembly to be under the control of the spindle assembly checkpoint (SAC) as SAC arrested cells maintain CDK1 mediated phosphorylation status of proteins. My data clearly demonstrates that this is the case. In the presence of benomyl, when the SAC is active when cells enter mitosis, the nucleolar disassembly of Bop1 is completely prevented while the nucleolar disassembly of Fibrillarin is only partially prevented. The reassembly of the dispersed nucleolar proteins, like Fibrillarin, in the presence of an activated SAC does not occur suggesting the removal of phosphate groups from proteins phosphorylated by CDK1 may be required for nucleolar reassembly. Secondarily, in the presence of an active SAC, the NOR may not be removed from the nucleolus possibly preventing complete disassembly and re-assembly of nucleolar proteins. This is supported by the data where benomyl treated cell unable to engage an active SAC (Mad2 deleted mutants) and which undergo CDK1 inactivation continue on with nucleolar disassembly and reassembly even in the absence of nuclear division. This strongly suggests that degradation of cyclin B, which in turn results in inactivation of CDK1, may

171 play a specific role in nucleolar disassembly and reassembly. Data with non-degradable cyclin B demonstrates that the inactivation of CDK1 is necessary for the disassembly of late processing proteins like Bop1 but is not necessary for the disassembly or reassembly of early proteins like Fibrillarin. It is thus possible that CDK1 mediated regulation of nucleolar proteins specifically directed at certain proteins (in this case Bop1) may be responsible for the step-wise disassembly of the parental nucleolus from the remnant. Such a regulation of disassembly of the nucleolus by CDK1 phosphorylation also demands that an opposing phosphatase play a role in nucleolar reassembly. The presence of the type I phosphates BIMG in the nucleolus suggests that such a potential role may be played by BIMG. BIMGs localization to the interphase nucleolus and then to the mitotic nuclear remnant suggests that BIMG may help dephosphorylated proteins in the remnant resulting in their disassembly from the remnant and allowing their reassembly into daughter nucleoli.

7.3.4. Why divide the A. nidulans nucleolus via a cytoplasmic intermediate as opposed to its segregation on the DNA? The nucleolus in the yeast systems is divided with DNA along the spindle during closed mitosis and serves as a much simpler mechanism of nucleolar segregation. Why is it that the A. nidulans nucleolus undergoes such a distinct type of nucleolar segregation? What is so different between A. nidulans and the yeast? The mitotic process in A. nidulans is relatively much shorter than in yeast. The average time taken for mitosis to finish in A. nidulans is only 5 to 7 minutes as opposed to a much longer mitosis in yeast. This quick mitosis is aided by the partial disassembly of the NPC which allows for the rapid influx of tubulin and thus the rapid assembly of the mitotic spindle. However, opening of the nuclear pores in this manner, though perhaps highly beneficial for mitotic entry, might cause problems for the nucleus to exit mitosis. For instance, the opening of the pores will allow many cytoplasmic proteins which normally lack a nuclear localization signal (NLS) to gain access during this process and localize in the nucleus. These proteins at the same time might not contain a nuclear export signal and hence pose a problem in their removal by active transport after cells finish mitosis. The

172 contamination of the nucleolus by these proteins is thus highly likely during the partially open mitosis of A. nidulans. A possible mechanism of getting rid of these unwanted proteins from the nucleolus would be to put the individual nucleolar components through a cycle of disassembly and reassembly from the cytoplasm into the newly formed nucleolus. In this manner only real nucleolar proteins belonging to the nucleolus would be able to get back into the nucleolus. An alternative mechanism of cleaning the nucleolus and getting rid of non-nucleolar proteins would be to release all its contents to the cytoplasm similar to NLS-DsRed and selectively re-import them back to the nuclei. However, as mentioned earlier, this may result in unequal distribution of proteins among neighboring nuclei undergoing mitosis at slightly different times as seen for NLA-DsRed. On the other hand, the nucleolus locally releases its content into the cytoplasm from a remnant structure in between the two newly forming daughter nuclei and coincide this release with the resumption of active nuclear import. This may ensure that nuclei undergoing mitosis at slightly different timings still contain equal amounts of nucleolar proteins as opposed to proteins dispersed in the cytoplasm like NLS-DsRed. This idea is supported by the mathematical modeling data presented. A summarized version of the entire nucleolar segregation process is shown in Figure 7.13. Mitosis is marked by the partial disassembly of the NPC at prophase during which the DNA condenses and the spindle pole bodies begin to separate (panel B). At metaphase, the spindles are assembled and DNA is poised to segregate along the mitotic spindle (panel C). The seclusion of the nucleolar proteins away from the chromatin such that it exists without the presence of the NOR during anaphase is shown in panel D. The NE then undergoes a doubly pinched such that two daughter nuclei are generated and the nuclear remnant containing the nucleolar proteins is formed (panel E). This occurs during late telophase/early G1. Following this, the dispersed nuclear pore proteins specifically return to the newly formed daughter nuclei in early G1 (panel F). The form of NE at this stage differs between the daughter nuclei and the nuclear remnant as the NE varies in their composition of nuclear pore proteins and their transport capability. The daughter nuclei contain all the nuclear pore proteins and are transport competent unlike the

173 remnant. Once nuclear transport resumes, the nucleolar proteins begin to disperse from the remnant into the cytoplasm and are concomitantly re-transported into the daughter nuclei (panel F, in yellow). Nucleolar proteins like Bop1 disperse later and are subsequently reassembled into daughter nuclei later (panel G, in green). By G1, the nucleolar proteins are all reassembled into daughter nuclei resulting in the formation of two functional nucleoli and fully functional nuclei (panel H).

174

G2 G2C G2

M M M

G1 G1 Gle1 - GFP NE DNA Merge

G1

Erg4 - GFP

(A. Image courtesy of Osmani A. H, B. & C. Image courtesy of Liu HL)

Figure 7.1. Generation of two daughter nuclei during mitosis in A. nidulans occurs by a double pinch of the nuclear membrane which forms a transient nuclear remnant devoid of DNA (A) An-Gle1-GFP remains at the NE throughout mitosis and clearly locates around three structures during late mitosis. The arrowhead marks the central transient structure (B) An-Erg4-GFP signal reveals the distribution of the ER and the NE during mitosis and clearly shows the existence of the three nuclear structures. The arrows mark the points of double pinch which gives rise to two daughter nuclei and a nuclear remnant (C) An-Erg4- GFP with H1-RFP DNA reveals the nuclear remnant (arrowhead) to be devoid of DNA.

175 B a

b

c

d

e

f

g

Figure 7.2. The nucleolus of A. nidulans segregates during mitosis via a mechanism that generates three nucleolar structures (A) Mitotic segregation of nucleolar protein An-Fib-GFP. The parental nucleolus (arrowhead, panel c) of A. nidulans undergoes disassembly at more or less the same time that new nucleoli are being reformed on either side. (B) An-Fib-GFP with H1-mRFP. As DNA is segregated during anaphase and telophase Fib-GFP does not segregate with the DNA (panel c). Instead the Fib-GFP mass is left behind, in between the separated DNA (arrowhead, panel d). After DNA is completely segregated new nucleoli begin to appear within each segregated mass of DNA (panel d). The appearance of the expanding daughter nucleoli corresponds with a proportional decrease in the parental Fib-GFP signal. With time the entire parental Fib-GFP signal is lost and all Fib-GFP becomes located within daughter nuclei. These G1 nuclei thus appear as smaller version of the starting G2 nuclei (panel g).

176

Figure 7.3. Mitotic segregation of GFP tagged nucleolar proteins CgrA, Bop1 and Nrap The pattern of segregation of CgrA-GFP, Bop1-GFP and Nrap-GFP in relationship to segregation of DNA was identical to that of Fib-GFP (Figure 7.2)

177

A

Erg4-GFP Fib-chRED Merge

B a C a

b b

c c

d d

e e TOPO1 Fib-chRED Merge TOPO1 H1-mRFP Merge

Figure 7.4. The transient nuclear remnant formed by double NE pinch contains the nucleolus but not the nuclear organizing region (NOR) (A) Erg4-GFP (ER marker) with Fib-chRED (nucleolus), reveal the nuclear remnant formed by double pinching of the NE membrane contains the nucleolus. (B) NOR marker TOPO1 with DNA, shows the NOR to segregate with DNA. (C) NOR marker TOPO1 with Fib-chRED, shows the nuclear remnant to be devoid of the NOR. Thus, the nucleolus in the nuclear remnant can exist in the absence of the NOR.

178

Figure 7.5. Daughter nuclei re-establish transport before reassembling nucleolar proteins NLS-DsRed with Fib-GFP shows NLS-DsRed to accumulate first in the nucleus (panel c, arrows) before Fib-GFP (panel g). This indicts that nuclear transport is first established and then new daughter nucleoli are formed in G1.

179 a

b

c

Fib-CR Bop1-GFP Merge

Figure 7.6. Fib-GFP with Bop1-chRED shows step-wise disassembly and reassembly The simultaneous visualization of Fib-GFP with Bop-chRED clearly establishes the timing of disassembly of Fib-GFP from the parental nucleolus and reassembly into daughter nucleoli (red bar)to be before that of Bop1-chRED (green bar).

180 Wild Type WT + Benomyl Mad2del + Benomyl A B C

*

Fib-GFP NLS-DsRed Merge Fib-GFP NLS-DsRed Merge Fib-GFP NLS-DsRed Merge

(Inset) Figure 7.7. The final disassembly of Fib-GFP is under the control of SAC (A) Disassembly and reassembly of Fib-GFP in wild- type nuclei during which NLS-DsRed is first dispersed and then returned to daughter nuclei. (B) Mitosis in the presence of Benomyl, NLS-DsRed remains dispersed for an extended period of time. Fib-GFP undergoes an initial disassembly but for the most part remains intact and does not undergo further disassembly. (C) Fib-GFP in a Mad2 strain in Benomyl. Cells proceed with mitosis without nuclear division and Fib- GFP is seen to undergo disassembly from the nucleolus (inset, arrowhead) and reassembly into a new spatially separated nucleolus (inset, arrow).

181 WT WT + Benomyl Mad2del + Benomyl

*

Bop1GFP NLS- DsRed Merge

Bop1-GFP NLS-DsRed Merge Bop1-GFP NLS-DsRed Merge (Inset)

Figure 7.8. The disassembly of Bop1-GFP is under the control of SAC (A) Disassembly and reassembly of Bop1-GFP in wild- type cell during which NLS-DsRed is first dispersed and then returned to daughter nuclei. (B) Mitosis in the presence of Benomyl, NLS-DsRed remains dispersed for an extended period of time and Bop1-GFP remains intact and does not undergo any disassembly. (C) Bop1-GFP in a ΔMad2 strain in Benomyl. Cells proceed with mitosis without nuclear division and Bop1-GFP is seen to undergo disassembly from the nucleolus (inset, arrowhead) and reassembly into a new spatially separated nucleolus (inset, arrow).

182

Control Bop1 + Bop1 + A BDsRe WT cyclin B non-degradable cyclin B Non-degradable cyclin B Cyclin B Total DNA amount Bop1 of Bop protein Merg Fibrillarin Telophase arrest (Tel) Metaphase arrest (Met) Time spent in mitosis after addition of Benomyl Fibrillarin + t=0, Non-degradable cyclin B Entry into mitosis C

DNA

Fib

Merge Telophase arrest (Tel)

Reforming daughter Nucleoli (R)

Figure 7.9. The role of cyclin B in nucleolar protein disassembly (A) Graph showing proteins levels of cyclin B, Bop1 and Fib in a SAC arrested cell. Bop1 and cyclin B levels are maintained through the SAC as deduced from previous data. Fib is shown to undergo an initial disassembly even in the presence of the SAC. (B) In the presence of non-degradable cyclin B, cells arrest at telophase with segregated DNA as shown by DAPI staining in blue. Bop1-GFP however remains intact between the two DNA masses with no apparent disassembly of the protein. Control wild type cells show the typical localization of Bop1-GFP offset from the DNA. (C) In the presence of non- degradable cyclin B expression, Fib-GFP undergoes similar disassembly as seen in a SAC arrested cell. In addition, Fib-GFP is also seen to reassemble into the daughter nucleoli, suggesting no role of CDK1-cyclin B in Fibrillarin disassembly.

183 Control Bop1 + Fibrillarin + Bop1 + WT cyclinB non-degradable cyclinB non-degradable cyclinB

Non- Mitotic Non- Mitotic Non- Mitotic mitotic mitotic mitotic

93 7 63 37 65 35

Tel Met R Tel Met R Tel Met R 2 5 0 25 12010 25 9

Table 7.1 Effect of non-degradable cyclin B on Bop1 and Fib disassembly The table quantifies the number of mitotic cells when non-degradable cyclin B is expressed vs. wild type cells with no such expression in two separate strains containing Bop1-GFP and Fib-GFP. Additionally, cells arrested in telophase (Tel) vs. metaphase (Met) are noted. The number of nuclei with reassembled Fib-GFP is noted under R. All telophase arrested Bop1-GFP cells had intact Bop1 signal while all telophase arrested Fib-GFP cells showed cytoplasmic signal for Fib-GFP. Of this 40% of the cells showed reassembled Fib-GFP into the daughter nuclei.

184 A B

a

b

BIMG - GFP c

BIMG Nucleolus Merge

Figure 7.10. BIMG localization and distribution during mitosis (A) BIMG localizes to several sub-cellular locations including the spindle pole, the nucleoplasm, the germ tip and the nucleolus. BIMG during mitosis localizes to the nuclear remnant and is in between the two segregating spindle poles. (B) BIMG with Fib- chRED, shows BIMG to localize to the nucleolus and kinetochores during mitosis (panel a). BIMG co-localizes with Fib-chRED during nuclear remnant formation (panel b) and finally re-distributes to both daughter nuclei at mitotic exit (panel c).

185

Figure 7.11. Comparative reassembly of NLS-DsRed and nucleolar protein Fib Semi-synchronous mitosis in A. nidulans sometimes results in the entry into mitosis of one nucleus (marked 1) earlier than the other (marked 2). Entry into mitosis is followed by dispersion of NLS-DsRed into the cytoplasm and exit is marked by its active re-import into the daughter nuclei. Note however that 1 enter and exits mitosis first and therefore re-accumulate NLS-DsRed before its neighboring nuclei ultimately resulting in unequal NLS-DsRed distribution. The nucleolar protein Fib-GFP amounts in all nuclei however are relatively equal. This most probably is due to localization of Fib-GFP into the remnant and its slow dispersion in the vicinity and in between newly reforming daughter nuclei.

186 A B CD

E FG H

Figure 7.12A. Mathematical modeling of a Ds-Red like protein that disperses in the cytoplasm on onset of mitosis and is reassembled from the cytoplasm (A) Total dispersed protein in cytoplasm is shown as a green curve. (B) Nucleus 1 (N1) exits mitosis and its daughter nuclei begin to reimport the cytoplasmic protein represented by the two rising blue peaks and a decrease in the total protein shown by the green curve. (C and D) N1 daughter nuclei continue to reimport. (E) Nucleus 2 (N2) exits mitosis and daughter nuclei of N2 begin to reimport represented as the other two blue peaks. (F) N2 daughter nuclei continue to reimport whatever is left of the cytoplasmic protein, note that N1 daughter nuclei are also continuing to reimport. (H) shows zero cytoplasmic protein and an end of reimport into nuclei. Note that N1 daughters have more protein in them compared to N2 daughters.

187 A B CD

E FG H

Figure 7.12B. Mathematical modeling of a protein restricted to the nuclear remnant during mitosis (A) shows two nuclei in mitosis that have secluded nucleolar proteins into a remnant like structure represented by the two red peaks. (B) Nucleus 1 (N1 on left) exits mitosis first and a concomitant release of remnant protein in the middle of the two daughter nuclei occurs (green curve) and its reimport into daughter nuclei can be seen as the two rising blue peaks. (D and E) The N1 remnant continues to disperse protein locally in the cytoplasm in vicinity of the two daughter nuclei of N1 which continue to reabsorb and this can be seen as a decrease in the red peak with proportionate increase in the two neighboring blue peaks. (F) Nucleus 2 (N2) exits mitosis and its remnant protein (red peak on right) begins to disperse protein into the cytoplasm at the same time as N2 daughter nuclei begin to reimport. (G) shows the time differential between the N1 remnant and N2 remnant however in (H) all four daughter nuclei have equal remnant protein showing that localized dispersal of protein in between reforming daughter nuclei lessens competition between nuclei and allows for more equal segregation of remnant like proteins.

188

Figure 7.13 Summarized model for mitotic segregation of the nucleolus in A. nidulans (A) A. nidulans nucleus with intact NE, duplicated spindle poles, NOR and the nucleolus assembled around the NOR. (B) At prophase, the NE undergoes partial disassembly if it’s NPC and opens the nuclear pores to the outside cytoplasm, resulting in equilibration of soluble proteins and RNA. (C) At metaphase, spindle formation occurs and the DNA is all set to segregate (D) DNA segregates during anaphase leaving behind the nucleolus. The NOR segregates with DNA. (E) Double pinch of the NE occurs in telophase resulting in two distinct forms of NE. The NE around the DNA is same as normal NE. The NE around the nuclear remnant is lacking certain Nups, is transport incompetent. The nuclear remnant is devoid of DNA and contains nucleolar proteins. (F) The NE around daughter nuclei reassembles dispersed Nups and restart nuclear transport. The nucleolar proteins disassemble into the cytoplasm and are re-imported before being reassembled at the NOR in the daughter nuclei reconstituting daughter nucleoli. (G) Step- wise disassembly and step-wise reassembly results in early processing protein Fib to appear first than late processing protein Bop1. (H) By G1, nucleolar reassembly is completed and two functional nuclei with respective functional nucleoli are made.

189

NOR Spindle NE A poles Nucleolus

H NPC disassembly

G2 B

prophase

G

Sequential dispersion metaphase and reassembly of the G1 nucleolus C F

early G1 anaphase

telophase/G1 E D Double pinch of NE around nucleolus

Figure 7.13

190

CHAPTER 8

FINAL DISCUSSION

8.1. Overview The aim of this project was to identify and characterize NIMA interacting proteins and proteins that may help regulate the function of NIMA in A. nidulans. Activation of the NIMA protein kinase is absolutely required for all aspects of mitosis in A. nidulans. I have used a copy number suppressor screen utilizing the AMA1 plasmid to screen for genes that suppress the nimA1 mutant phenotype when expressed in excess. Three genes encoding potential NIMA interacting/regulatory proteins were identified in this screen, mcnA, mcnB and mcnC. Overexpression of each of these genes had specific effects on NIMA protein levels and/or its potential substrates. Mitosis in all organisms involves a massive reorganization of cellular structure. The microtubule cytoskeleton is organized into mitotic spindles between the duplicated spindle pole bodies accompanied with chromosomal condensation followed by attachments of these chromosomes via their kinetochore to the spindles. With the coordinated effort of multiple motors the chromosomes are then segregated into two daughter nuclei. Mitosis in all organisms also sees a substantial wave of protein phosphorylations, controlling signal events that coordinate mitotic processes and ensure accurate mitotic entry. One of the key switches for the onset of mitosis is NIMA. NIMA, a serine/threonine kinase, is one of a few select protein kinases proven to be essential for the regulation of different aspects of mitosis. No wonder then that its regulation through the cell cycle is highly regulated.

191 The understanding of nimA regulation prior to the screen includes information regarding the following. The NIMA protein and mRNA levels are regulated such that they accumulate in late G2, reach their peak at M and are down-regulated for mitotic exit. The down-regulation of the protein level is known to occur through PEST sequence mediated protein degradation while the kinase activity of the protein is regulated via specific phosphorylation events resulting in hyper-phosphorylation of the protein. The sub-cellular localization of NIMA also plays a role in determining the substrates of the kinase and acts as another level of regulation. In spite of knowing many aspects of nimA regulation, certain information regarding its transcriptional control, post-transcriptional control, and its positive regulators has been largely unknown. The advantage of performing the copy number screen to rescue a mutant phenotype of nimA was the identification of proteins that may interact with nimA and the identification of additional levels at which nimA is regulated. Note that the conditional nimA1 mutation results in a G2 arrest phenotype due to the inability to locate NIMA1 protein to its proper sub- cellular region and not because of lack of kinase activity as the NIMA1 protein contains partial kinase activity. Surprisingly, all three copy number suppressors only partially compensated for the nimA1 growth defect suggesting suppression of nimA1 might involve the modulation of NIMA1 protein itself.

8.2. mcnB probably transcriptionally activates nimA Over expression of mcnB specifically results in the up-regulation of NIMA protein levels. The presence of a forkhead domain found in other known transcription factors clearly suggests MCNB to be a transcription factor. Additionally, study of related homologs from yeast have shown them to be involved specifically in the regulation of gene expression during the G2-M transition, the same window of time during which NIMA performs its functions. The regulation of nimA by mcnB may be direct transcriptional activation or may involve the activation of another protein that in turn may up-regulate NIMA levels. The loss of mcnB function had no obvious effect on cellular function strongly suggesting a redundancy for this proteins function. Importantly, there are four related MCNB like proteins which exist in A. nidulans and all contain the

192 forkhead domain. One or more of these MCNB related proteins could thus play a redundant role with MCNB which would explain the lack of cell cycle defects in the absence of MCNB function. More experiments need to be done to define if mcnB directly activates nimA transcription and if the other related proteins also act as redundant nimA specific transcription factors.

8.3. mcnC might regulate NIMA sub-cellular localization mcnC gene over expression did not affect NIMA1 levels suggesting an alternative mechanism for nimA1 mutant suppression. An earlier extragenic suppressor screen identified two potential nimA substrates, SONA and SONB, both nuclear pore proteins that are thought to be phosphorylated by NIMA at mitosis to promote opening of the nuclear pore resulting in NIMA entry into the nucleus. Similarly, when over expressed, mcnC caused dispersion of SONB from the nuclear pores. This might then “open” the nuclear pore to the extent that NIMA1 might be able to gain access to its nuclear substrates and so promote mitosis. This mechanism might explain how extra MCNC suppresses nimA1 without overtly affecting its protein expression levels. mcnC behaves like a positive regulator of nimA and consistent with this possibility its deletion in combination with two other temperature sensitive alleles of nimA causes synthetic lethality. One unexpected and surprising effect of MCNA over expression was on cell morphology and spore germination. However, recent work on the location of the NIMA kinase might shed light on this phenomenon. In addition to localizing to nuclei, and mitotic spindle structures, NIMA has also been seen to locate to the growing tip of cells and also to the sites of cytokinesis, which in A. nidulans are the septa. These additional locations of NIMA have only recently been documented and suggest NIMA plays roles in cell polarization and septation. mcnC over-expression, which results in specific defects in polarization and cell tip emergence, therefore suggests that mcnC may regulate these function of nimA as well.

193 8.4 mcnA is a novel nucleolar associated protein The sub cellular localization of mcnA and its co-localization to the nucleolus through most of interphase suggests that it is a novel nucleolar protein. However, its cell cycle behavior differs from other unknown nucleolar proteins in terms of its mitotic segregation. The regulated localization of MCNA protein to the cytoplasm at M-G1 transition suggests a specific role for MCNA in M and G1. Its resemblance to some features of TAM bodies hint at its function being in mRNA or protein degradation and may prove to be an additional step to down regulate NIMA proteins/mRNA levels during mitotic exit resulting in a rapid shut down of NIMA function. The non-essentiality of mcnA function suggests that this function most probably aids in a process that already exists for NIMA protein degradation, for example the APC mediated protein degradation pathway. Importantly the localization of MCNA to the nucleolus may serve to prevent its function during the period of the cell cycle when not required. This is analogous to the known regulation of the Cdc14 phosphatase which plays a role during yeast exit from mitosis. Cdc14 is kept inactive in part by being sequestered in the nucleolus during interphase. Additionally, MCNA may serve a completely different and novel function at the nucleolus itself and in fact MCNA defines a new sub-nucleolar compartment. This study therefore identified additional levels of regulation of nimA expression (both up-regulation and down regulation) and NIMA localization. As mentioned before, the suppression of nimA1 mutant cells is a partial recovery of their growth defect at the restrictive condition and not a full restoration. This suggests that the mode of suppression by these genes might involve modulation of the partially active NIMA1 kinase expression and localization. The copy number suppressor thus identified genes that probably directly affect the functioning of nimA and show the usefulness of such a screen. Additionally future studies of physically interacting proteins through pull down studies using the S- tagging system will allow us to expand our understanding of other players that may be involved in the nimA regulation pathway.

194 8.5. A new mechanism for mitotic segregation of the nucleolus The study of the nucleolus was spearheaded due to the identification of MCNA which localized to the nucleolus and segregated during mitosis in a very unique pattern. We were thus curious to know if all nucleolar proteins in A. nidulans segregated during mitosis in a similar pattern in which it is left outside of the nucleus in the cytoplasm during chromosomal separation and is segregated to daughter nuclei in G1. To our surprise, other nucleolar proteins also divided during G1 after chromosomal segregation on the mitotic spindle. The data clearly shows that unlike yeast, the nucleolus in A. nidulans is disassembled in the cytoplasm during late mitosis and reassembled during G1 to form daughter nuclei. The timing of reassembly of MCNA was found to extend beyond that of the other nucleolar proteins studied which may mean that MCNA is not a bonafide nucleolar protein. The specific disassembly and reassembly of nucleolar proteins resembles the mitotic segregation of the nucleolus in higher eukaryotes. In higher eukaryotes undergoing an open mitosis with NE breakdown, the nucleolar components move away from the nucleolar organizing region (NOR) and locate to the perichromosomal area from where they are specifically recruited back in a particular order to the new daughter NORs at G1. Highly similar to this in A. nidulans, where mitosis is partially open with an intact NE but open nuclear pores, the nucleolus is specifically removed from the NOR at mitotic exit and expelled to the cytoplasm from where it is reassembled in a step wise manner to the newly forming daughter nucleoli. The step-wise reassembly is thought to follow the order in which the proteins are required for ribosomal related activity in the new nucleolus. This ordered reassembly is identical for different proteins between A. nidulans and higher eukaryotes suggesting that the mitotic segregation mechanisms may well be conserved among them. This study helped us gain important insights into the fundamental steps involved in mitotic segregation of the nucleolus in A. nidulans. However as is with any new discovery, the number of questions that remain unanswered exceed if not equal the number of answers found. For example, we now know that the NE surrounding the two newly forming daughter nuclei differs in form compared to the NE surrounding the nuclear remnant structure. What is it though that regulates and maintains this difference

195 in the NE, allowing only the daughter nuclei to become functional and transport competent entities while making the remnant a transient structure? Similarly, the double pinch physically separates the bulk of the nucleolar proteins from the rest of the DNA but the possible motors involved in this step remain unknown. We also do not have a solid idea regarding what may regulate these motors and the exact location and timing of the pinches. The nuclear remnant containing the nucleolar proteins also need regulation in order to have a timely step-wise dispersion of its proteins to the cytoplasm such that they can be reassembled in a particular order to the newly forming daughter nucleoli. What then may keep the remnant structure intact in the cytoplasm with its nucleolar content within till it is time to disassemble? My findings suggest that inactivation of cyclin B likely plays a key role but this is unlikely to explain all aspects of nucleolar disassembly- reassembly. Thus while the mechanics of nucleolar segregation in A. nidulans is a completely novel discovery, many key questions regarding its regulation and execution now remain to be answered. The regulation of various aspects of mitosis by NIMA kinase such that it occurs properly is absolutely essential in A. nidulans and probably certain aspects of nucleolar segregation may also be under regulation by NIMA. Although the function of nimA in organisms like Saccharomyces cerevisiae does not seem to be as important during mitosis as it is in A. nidulans, this may be explained by the stark differences in mitosis between the two organisms. Firstly, S. cerevisiae does not undergo any nuclear pore complex disassembly and performs mitosis within an intact closed nuclear envelope. Secondly, it segregates its nucleolar proteins at the same it segregates its DNA. Unlike S. cerevisiae, A. nidulans undergoes partial disassembly of both its nuclear pores and nucleolar proteins and thus likely requires additional levels of regulation for each step. This may be one of the reasons why NIMA is indispensable for mitosis in A. nidulans. Further support for this idea comes from studies of the NIMA protein kinase family in higher eukaryotes like humans. Mitosis in humans not only involves disassembly of the nuclear pores but also the complete break down of the NE in addition to the complete disassembly of the nucleolus. There are at least 11 different nimA related kinases with varied mitotic and non-mitotic functions in humans. This expansion of the nimA gene family, from one in A.

196 nidulans to eleven in humans, may reflect the more complex mitotic processes of higher eukaryotic open mitoses. Clearly the fact that A. nidulans undergoes both nuclear pore and nucleolar disassembly-reassembly during its mitosis makes it an ideal model genetic system in which to further define the molecular mechanisms regulating these most fascinating aspects of the miracle that is mitosis.

197

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