CHARACTERIZATION OF TWO NIMA INTERACTING SUGGESTS

A LINK BETWEEN NIMA AND NUCLEAR MEMBRANE FISSION

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of

Philosophy in the Graduate School of The Ohio State University

By

Jonathan Robert Davies, M.S.

* * * * *

The Ohio State University

2004

Dissertation Committee: Approved by:

Dr. Stephen A. Osmani, Adviser

Dr. Lee F. Johnson ______Adviser Dr. Berl R. Oakley Department of Molecular Genetics Dr. Paul K. Herman

ABSTRACT

In the filamentous fungus , the NIMA is required along with CDK1/cyclinB for mitotic entry. The essential function of

NIMA in A. nidulans and the growing recognition of its importance in other eukaryotes, means that the study of NIMA function should reveal unique insights into regulation amongst a broad range of organisms. I describe here the characterization of TINC and TIND, two NIMA interacting proteins identified in a yeast Two-hybrid screen, and describe the potential novel roles they may play in mitotic regulation.

TINC and a related in A. nidulans, An-HETC, are highly similar to proteins conserved in filamentous fungi. Strains which lack both tinC and An- hetC are viable, but do display osmotic and cold sensitivity.

Characterization of TINC suggests that it is involved in mitotic regulation.

First, TINC is present in the nucleus during . Second, TINC interacts with

NIMA in a phosphorylation state dependant manner. Third, truncated forms of

TINC (∆N-TINC) produce cell cycle defects characterized by a defect in nuclear membrane fission in which cells are able to separate DNA but unable to cleave the nuclear envelope. Significantly, ∆N-TINC localizes to membranous bodies

ii which associate with nuclei. Finally, expression of ∆N-TINC promotes premature loss of NIMA from mitotic samples.

The second NIMA interacting protein, TIND, is conserved from bacteria through to humans. TIND is predominantly mitochondrial throughout the cell cycle, but work in other organisms suggests that alternate forms of TIND may exist outside the mitochondria. Expression of a form of TIND which lacks the mitochondrial targeting peptide (∆N-TIND) produces nuclear division and nuclear membrane fission defects similar to those seen for ∆N-TINC. Additionally, ∆N-

TIND also produces defective mitotic spindles, which appear monopolar, an effect not seen for ∆N-TINC.

The facts that TINC and TIND were isolated as NIMA interacting proteins,

TINC interacts with NIMA in A. nidulans, and expression of ∆N-TINC or ∆N-TIND produces specific defects in nuclear membrane fission suggest roles for these proteins in mitotic regulation. Additionally, the finding that NIMA is destabilized in cells displaying mitotic defects suggests a role for NIMA in regulating nuclear membrane fission.

iii

DEDICATION

This work is dedicated to my parents,

to my wife Heather,

and to my son Colin.

iv ACKNOWLEDGMENTS

I would like to thank my adviser, Dr. Stephen Osmani, for his guidance support, and friendship throughout my graduate studies. The first summer I spent in Steve’s lab as an undergraduate provided the motivation for me to pursue a graduate degree in the Life Sciences.

I would like to thank the members of my committee, Dr. Lee Johnson, Dr.

Berl Oakley, and Dr. Paul Herman for their time and guidance during my graduate studies at The Ohio State University.

I would also like to acknowledge the current and past members of the

Osmani Lab for their friendship and guidance. Specifically, I wish to thank Aysha

Osmani for taking the time to teach me the techniques needed to work in the lab.

I also wish to thank Dr. Colin De Souza for his willingness to talk with me about my project and for his generosity in providing strains.

This research was funded by a grant from the National Institutes of Health.

v VITA

1997………………………………………...B.S. Biology, Grove City College

1999………………………………………...M.S. Physiology, The Pennsylvania State University College of Medicine

1999 - 2000………………………………..Graduate Research Associate, The Pennsylvania State University College of Medicine

2000 – present…………………………….Graduate Research Associate, The Ohio State University

PUBLICATIONS

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

Osmani, A.H., Davies, J., Oakley, C.E., Oakley, B.R., Osmani, S.A. (2003). TINA interacts with the NIMA kinase in Aspergillus nidulans and negatively regulates astral microtubules during metaphase arrest. Mol. Biol. Cell. 14, 3169-79.

Miller, B.A., Zhang, M.Y., Gocke, C.D., De Souza, C., Osmani, A.H., Lynch, C., Davies, J., Bell, L., Osmani, S.A. (1999). A homolog of the fungal nuclear migration nudC is involved in normal and malignant human hematopoiesis. Exp. Hematol. 27, 742-50.

FIELDS OF STUDY

Major Field: Molecular Genetics

vi TABLE OF CONTENTS

Page

ABSTRACT...... ii

DEDICATION ...... iv

ACKNOWLEDGMENTS ...... v

VITA ...... vi

LIST OF FIGURES...... xii

LIST OF TABLES ...... xiv

CHAPTER 1. INTRODUCTION...... 1

1.1. PROBLEM STATEMENT...... 1 1.2. ASPERGILLUS NIDULANS ...... 2 1.2.1. General classification and description...... 2 1.2.2. Asexual life cycle ...... 3 1.2.3. Sexual life cycle ...... 4 1.2.4. A model for cell cycle research ...... 5 1.3. THE CELL CYCLE ...... 9 1.3.1. Overview...... 9 1.3.2. Interphase...... 10 1.3.3. M Phase...... 11 1.3.4. Regulation...... 13 1.3.4.1. 2001 Nobel Prize: ...... 13 1.3.4.2. Checkpoints:...... 13 1.3.4.3. Identification of cyclin dependant kinases: ...... 16 1.4. NIMA PROTEIN KINASE...... 19 1.4.1. Isolation ...... 20 1.4.2. Functional regions within NIMA ...... 20 1.4.3. NIMA fluctuations through the cell cycle ...... 24 1.4.4. Phosphorylation and NIMA activation ...... 25 1.4.5. Active NIMA is required for the G2-M transition...... 26 vii 1.4.6. NIMA interacting proteins and targets...... 27 1.4.7. NIMA related kinases and cell cycle control...... 28 1.5. AIMS...... 31

CHAPTER 2. MATERIALS AND METHODS...... 43

2.1. GENERAL DNA PREPARATION AND CLONING ...... 43 2.1.1. Plasmid maxiprep and miniprep...... 43 2.1.2. DNA cloning...... 44 2.1.3. Polymerase Chain Reaction (PCR)...... 44 2.1.4. Primers ...... 45 2.1.5. Automated fluorescent sequencing...... 45 2.1.6. Bacterial strains ...... 46 2.1.7. Transformation of bacteria ...... 46 2.1.8. Storage and stock preparation of bacteria ...... 47 2.2. CULTURE AND GENETICS OF A. NIDULANS ...... 47 2.2.1. A. nidulans specific media ...... 47 2.2.2. A. nidulans Strains...... 49 2.2.3. Preparation of A. nidulans conidia stock suspensions ...... 49 2.2.4. Conidiospore Quantitation ...... 50 2.2.5. Long term storage and stock preparation of A. nidulans...... 50 2.2.6. Strain generation by meiotic crossing ...... 51 2.2.7. Diploid formation...... 52 2.3. GENERAL A. NIDULANS TECHNIQUES...... 52 2.3.1. Small scale protein preparation ...... 52 2.3.2. Large scale protein preparation ...... 53 2.3.3. Small scale genomic DNA extraction...... 55 2.3.4. Large scale genomic DNA extraction...... 55 2.3.5. Transformation of A. nidulans ...... 56 2.3.6. Immunofluorescence...... 58 2.3.7. Staining of mitochondria ...... 59 2.4. ISOLATION OF FULL-LENGTH CDNAS BY 5’RACE PCR ...... 59 2.5. ISOLATION OF GENOMIC CLONES FOR TINC AND TIND...... 61 2.6. TINC ANTIBODY...... 62 2.7. HA TAGGING OF TINC AND TIND ...... 63 2.8. GFP TAGGING OF TINC AND TIND...... 64 2.9. ALCA DRIVEN PROTEIN EXPRESSION IN A. NIDULANS...... 64 2.10. EXAMINATION OF TINC AND TIND LOCALIZATION ...... 65 2.11. DELETION OF TINC AND AN-HETC...... 66 2.12. ∆TINC AND ∆AN-HETC PHENOTYPE TESTING...... 69

viii 2.13. CROSSES BETWEEN ∆TINC / ∆AN-HETC AND CELL CYCLE MUTANTS...... 70 2.14. EXAMINATION OF NUCLEAR DIVISION DEFECTS IN ∆N-TINC OR ∆N-TIND EXPRESSING CELLS ...... 70 2.15. EXAMINATION OF NUCLEAR ENVELOPES AND NUCLEAR MEMBRANE FISSION DEFECTS IN CELLS EXPRESSING ∆N-TINC OR ∆N-TIND ...... 71 2.16. EXAMINATION OF ∆N-TINC AND ∆N-TIND LOCALIZATION ...... 71 2.17. TINC AND NIMA CO IMMUNOPRECIPITATIONS ...... 72 2.18. EXAMINATION OF NIMA IN ∆N-TIND AND ∆N-TIND EXPRESSING CELLS ..... 73 2.19. MICROSCOPY AND IMAGE CAPTURE SOFTWARE...... 74 2.20. BIOINFOMATICS AND DNA ANALYSIS...... 75

CHAPTER 3. SEQUENCE CHARACTERIZATION AND SUBCELLULAR LOCALIZATION OF TINC AND TIND ...... 80

3.1. INTRODUCTION ...... 80 3.1.1. The yeast Two-hybrid system...... 81 3.1.2. NIMA interacting protein screen...... 82 3.1.3. Expression proteins in A. nidulans using the alcA promoter...... 84 3.1.4. Heterokaryon incompatibility...... 85 3.1.5. Fe-S clusters...... 87 3.2. RESULTS...... 88 3.2.1. tinC cloning and sequence characterization...... 88 3.2.2. TINC is a member of a fungal specific family of proteins ...... 90 3.2.3. TINC antibody characterization...... 92 3.2.4. TINC localization...... 93 3.2.5. tinD cloning and sequence characterization ...... 95 3.2.6. TIND is a highly conserved protein ...... 96 3.2.7. TIND localization...... 97 3.2.8. Identification of human tinD cDNAs ...... 98 3.3. DISCUSSION ...... 98 3.3.1. TINC and heterokaryon incompatibility ...... 98 3.3.2. TIND and Fe-S cluster assembly ...... 99 3.3.3. TINC localization...... 100 3.3.4. TIND localization...... 101

CHAPTER 4. TINC / An-HETC FUNCTION IS NON-ESSENTIAL ...... 119

4.1. INTRODUCTION ...... 119 4.1.1. A. nidulans transformation characteristics...... 119 4.1.2. A method for generating deletion targeting constructs...... 120

ix 4.2. RESULTS...... 121 4.2.1. Generation of ∆tinC, ∆An-hetC, ∆tinC/∆An-hetC strains ...... 121 4.2.2. Deletion phenotype testing ...... 122 4.3. DISCUSSION ...... 124

CHAPTER 5. TINC, TIND, AND NUCLEAR MEMBRANE FISSION ...... 128

5.1. INTRODUCTION ...... 128 5.1.1. Closed mitosis...... 129 5.1.2. Nuclear membrane fission ...... 131 5.1.3. Examination of the nuclear envelope in A. nidulans...... 132 5.2. RESULTS...... 133 5.2.1. ∆N-TINC expression but not TINC expression produces nuclear division defects...... 133 5.2.2. ∆N-TINC is present in membranes ...... 135 5.2.3. ∆N-TIND expression but not TIND expression produces nuclear division defects and abnormal mitotic spindles ...... 136 5.2.4. ∆N-TIND is present throughout the cell ...... 138 5.2.5. Cells expressing ∆N-TINC or ∆N-TIND proceed through faulty mitoses...... 138 5.2.6. Cells expressing ∆N-TINC or ∆N-TIND display nuclear membrane fission defects ...... 140 5.3. DISCUSSION ...... 142 5.3.1. TINC, TIND and growth inhibition ...... 142 5.3.2. Cell cycle specific defects ...... 142 5.3.3. ∆N-TINC and ∆N-TIND Localization...... 145

CHAPTER 6. TINC INTERACTS WITH NIMA IN A. NIDULANS...... 160

6.1. INTRODUCTION ...... 160 6.2. RESULTS...... 162 6.2.1. TINC interacts with NIMA in A. nidulans ...... 162 6.2.2. Expression of ∆N-TINC produces NIMA instability ...... 164 6.3. DISCUSSION ...... 166 6.3.1. TINC-NIMA Interaction...... 166 6.3.2. Premature loss of NIMA...... 167 6.3.3. A TIND-NIMA interaction in A. nidulans? ...... 167

x CHAPTER 7. FINAL DISCUSSION AND FUTURE DIRECTIONS ...... 172

7.1 OVERVIEW ...... 172 7.2. A POTENTIAL ROLE FOR TINC IN CELL CYCLE REGULATION ...... 174 7.3. A SEARCH FOR NEW ALLELES OF NIMA ...... 176 7.4. TWO MODELS ADDRESSING ∆N-TINC AND ∆N-TIND DEFECTS...... 177 7.5. ∆N- TIND AND REDOX REGULATION ...... 180 7.6. NIMA: MAINTAINING THE MITOTIC STATE AND NUCLEAR MEMBRANE FISSION .181

BIBLIOGRAPHY ...... 184

xi LIST OF FIGURES

Figure Page

Figure 1.1. Lifecycles of A. nidulans...... 33

Figure 1.2. Asexual life cycle 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 3.1. Identification of NIMA interating proteins using the yeast Two-hybrid system...... 105

Figure 3.2. pAL5...... 106

Figure 3.3. TINC...... 107

Figure 3.4. TINC is a member of a fungal-specific family of proteins...... 108

Figure 3.5. Characterization of a TINC specific antibody...... 110

Figure 3.6. TINC localization ...... 111

Figure 3.7 TIND ...... 113

Figure 3.8. TIND is a highly conserved protein...... 114

Figure 3.9. TIND localization ...... 115

xii Figure 3.10. TIND is mitochondrial during interphase and mitosis...... 117

Figure 3.11. Isolation of human tinD cDNAs...... 118

Figure 4.1 Deletion of ∆tinC and ∆An-hetC ...... 126

Figure 4.2. ∆tinC cells are osmotically sensitive and cold sensitive ...... 127

Figure 5.1. Nuclear membrane fission...... 148

Figure 5.2. Expression of ∆N-TINC inhibits growth in A. nidulans...... 149

Figure 5.3. Expression of ∆N-TINC produces nuclear division defects ...... 150

Figure 5.4. ∆N-TINC locates to the periphery of membranes...... 151

Figure 5.5. Expression of ∆N-TIND or TIND inhibits growth in A. nidulans ...... 153

Figure 5.6. Expression of ∆N-TIND produces nuclear division defects ...... 154

Figure 5.7. Expression of ∆N-TIND produces defective spindles ...... 155

Figure 5.8. ∆N-TIND locates throughout the cell ...... 156

Figure 5.9. Cells expressing ∆N-TINC or ∆N-TIND proceed through faulty mitoses ...... 157

Figure 5.10. Expression of ∆N-TINC or ∆N-TIND produces nuclear membrane fission defects ...... 158

Figure 6.1. TINC binds NIMA in A. nidulans ...... 169

Figure 6.2. Cells expressing ∆N-TINC prematurely lose NIMA ...... 171

Figure 7.1. Control of nuclear membrane fission and mitotic exit by NIMA ...... 182

xiii LIST OF TABLES

Table Page

Table 1.1. List of A. nidulans mitotic mutants ...... 42

Table 2.1. List of primers used for cloning and tagging protocols...... 76

Table 2.2. List of A. nidulans strains...... 78

xiv

CHAPTER 1

INTRODUCTION

1.1. Problem Statement

In growing and dividing cells, the ability to maintain and transmit a complete complement of genetic material to daughter cells is absolutely essential.

Misregulation of this process results in the production of disease states including cancer. Cellular growth and division are essential processes which are coordinated in a highly ordered continuum called the cell cycle. The basic patterns and mechanisms of cell growth and division are highly conserved from relatively simple eukaryotes including yeast and fungi through to higher eukaryotic systems such as humans. The regulatory elements which govern cell cycle progression are also highly conserved.

The cyclin dependant family of kinases is firmly established as one of the major regulators of the eukaryotic cell cycle. CDK1 is a central regulator of cell cycle progression in organisms from fungi to humans and is required for mitosis.

However, another essential protein kinase, referred to as NIMA, has been identified in the filamentous fungus A. nidulans. NIMA is a cell cycle regulated protein kinase which, along with CDK1, is required for the passage of cells into

1 mitosis in A. nidulans (Oakley and Morris, 1983; Bergen et al., 1984; Osmani et

al., 1987; Osmani et al., 1991a).

Present data confirm that CDK1 and NIMA are essential cell cycle

regulatory proteins in A. nidulans. Also, the identification of numerous NIMA related kinases in organisms ranging from yeast to humans, some with mitotic functions, suggest that a NIMA-like pathway of cell cycle regulation may exist in a wide range of organisms. The identification of NIMA interacting proteins and targets has already provided valuable insights into understanding how NIMA regulates the cell cycle. The identification and characterization of additional

NIMA interacting proteins will provide valuable insight into the role of NIMA in the fungal cell cycle and potentially that of higher eukaryotes. Overall, gaining a better understanding of cell cycle regulation in a genetically amenable model system such as A. nidulans will provide potentially valuable insight into the cell growth and division cycles of higher eukaryotes.

1.2. Aspergillus nidulans

1.2.1. General classification and description

Aspergillus nidulans is a filamentous ascomycete fungus first described in detail by Eidam in 1883 (Eidam, 1883). Aspergillus occurs ubiquitously in soil and plant matter, with spores present in air and water samples. The genus is very well characterized due to its varied roles within the pharmaceutical industry, human disease, and as a powerful model genetic organism. The citric acid

2 fermentation cycle of Aspergillus niger is important in the food industry and in

production of a number of pharmaceuticals, while Aspergillus fumigatus and

Aspergillus flavus are largely responsible for a range of respiratory based fungal infections broadly referred to as Aspergillosis. The experimental system employed in the work described in this thesis is A. nidulans. Over the past several decades, A. nidulans has become established as a powerful tool for genetic based research, since the first thorough genetic description of the organism by Pontecorvo in 1953 (Pontecorvo, 1953).

A. nidulans is capable of both asexual and sexual lifecycles (Figure 1.1).

While all the cell biological and biochemical experiments described here are performed using organisms undergoing asexual development, the sexual cycle was used to great extent for genetic manipulations.

1.2.2. Asexual life cycle

The asexual life cycle of A. nidulans is initiated by germination of a haploid uninucleate conidiospore (Figure 1.2A). Under favorable environmental conditions the spore undergoes polarized germination and rounds of synchronous mitoses resulting in the production of a multi-nucleate germling

(Figure 1.2B). Following the third or fourth mitotic division, the cell is physically compartmentalized by the production of a septum. Nuclei within this septal compartment cease further division while nuclei distal to the septum continue to divide. In liquid media, A. nidulans will continue to grow as a system of branched

hyphae, but fails to conidiate without an air interface. On solid media A. nidulans

3 is typified by radial growth of a branched hyphal network. Additionally, after a

period of growth specialized cells, termed foot cells, develop within the hyphae

(Figure 1.2C). These cells renter the cell cycle and produce the conidiophore,

the asexual reproductive structure of A. nidulans (Figure 1.2C).

The conidiophore consists of a stalk which supports a bulb shaped vesicle

(Figure 1.2C). Primary and secondary sterigma emerge from the vesicle (Figure

1.2C). The distal nuclei within each secondary sterigmata undergo rounds of

mitotic division with one nucleus remaining within the sterigmata while the other

nuclei becomes compartmentalized and develops into a conidiospore. This

process continues until each sterigma supports chains of conidia which may

number up to 100. These chains of conidia are responsible for the visible color

and general appearance of A. nidulans colonies. The color of conidiospores from

wild type isolates is green, but mutants are broadly available with varying spore

colors including chartreuse, yellow, fawn, and white (Figure 1.3). These color

mutations provide useful markers for genetic manipulations.

1.2.3. Sexual life cycle

In addition to an asexual cycle, A. nidulans is also capable of a sexual

cycle. A. nidulans is a self-fertile, homothallic, ascomycete. Unlike heterothallic

organisms, including the filamentous fungus Neurospora crassa, no mating types

exist within A. nidulans. The homothallic nature of A. nidulans has proved to be an advantage for genetic analysis due to the ability of the organism to undergo hyphal anastomoses and karyogamy. The sexual life cycle consists of three

4 basic steps: hyphal fusion to produce a heterokaryon, nuclear fusion to produce

a diploid, and meiosis resulting in haploidization (Figure 1.1). In practical terms,

two organisms with different nutritional requirements can be induced to undergo

hyphal fusion to form a heterokaryon on nutritionally limiting defined media. A

heterokaryon is an organism which contains mixed nuclei within a common

. While unstable, these heterokaryons can be maintained in a

laboratory setting. Additionally, at some frequency these nuclei undergo nuclear

fusion (karyogamy) to produce diploid nuclei. These diploids are stable, but can

be induced to haploidize in the presence of microtubule poisons. This

haploidization process is termed the parasexual cycle and has been used to

great effect to perform linkage analysis within the organism.

Alternately, in sexual development two nuclei undergo karyogamy, within

specialized cells termed asci to form a transient diploid. Meiotic division results

in the production of eight haploid ascospores per ascus (Figure 1.1). The

products of each meiosis are present within a single ascus, which allows for

tetrad analysis. Asci are contained within larger structures termed cleistothecium.

Cleistothecia may contain as many as 105 meiotic progeny, which provide an enormous number of “offspring” for genetic analysis.

1.2.4. A model for cell cycle research

In 1945, A. nidulans was first identified as a potentially valuable

"microorganism suitable for a genetic approach to certain problems of spatial organization of the cell" by geneticists at the University of Glasgow (Pontecorvo,

5 1945; Champe and Simon, 1992). Today, A. nidulans is recognized as a valuable model for study in a number of fields including cell cycle research.

Several aspects of the A. nidulans life cycle and genetics make it

particularly amenable to cell cycle research. A. nidulans has a relatively short

cell cycle time of ninety to 120 minutes, rapid growth rate, and well developed

genetics which include a large number of mutations which affect aspects of cell

cycle progression (Bergen and Morris, 1983). . When grown on rich media A.

nidulans has a cycle time of about 100 minutes consisting of a 15 minute G1 phase, a 40 minute S phase, a 40 minute and a 5 minute M (Bergen and Morris, 1983). Conidiospores can be stored in a dormant state in

suspensions for long periods. Following inoculation into media, the "awakened"

conidiospores provide a convenient and accurate initiation time for experiments

(Champe and Simon, 1992). Precise start points are particularly important in many cell cycle progression experiments. Additionally, as Aspergillus germinates rounds of synchronous nuclear division accompany apical hyphal growth. As a germ tube extends nuclei divide and migrate along its length (Figure 1.4). Within a single septal compartment all mitotic divisions occur nearly simultaneously, beginning at the apical portions of each hyphae and moving along its length in a rapid wave (Rosenberger and Kessel, 1967; Rosenberger and Kessel, 1968).

Due to this fact, it is easy to determine the number of nuclear division events which have occurred in a specified region of hyphae by simply counting the number of nuclei present. The generation of mutations in cell cycle regulatory frequently results in lethality necessitating the isolation of temperature

6 sensitive mutants. The ability of A. nidulans to grow over a broad range of

temperatures from 15°C to 44°C allows great opportunity for the isolation of

temperature sensitive and cold sensitive mutants (Doonan, 1992). The ability to

introduce transformed DNA through site-specific integration into the recipient

genome allows for complementation of mutant phenotypes and the ability to form

stable diploids and heterokaryons allows for dominant recessive

complementation analysis. Additionally, the lack of mating types and the

availability of a range of conidial color and auxotrophic mutants allows for the

generation of strains through meiotic recombination.

These characteristics coupled with the fact that earlier studies had demonstrated that A. nidulans exhibits the hallmarks of eukaryotic mitosis

(Robinow and Caten, 1969), prompted Ron Morris to choose A. nidulans as the model system in which he would perform a temperature sensitive screen to identify mutants involved in cell cycle progression (Morris, 1976). Genetic screens to identify cell cycle regulatory components were being conducted roughly at this same time by Paul Nurse and Lee Hartwell in S. pombe and S.

cerevisiae respectively (Hartwell et al., 1974; Nurse et al., 1976). Morris’s screen

was distinguished by the fact that the mutants were classified on the basis of

direct observations of nuclear and spindle morphology using techniques

described by Robinow and Caten rather than cell size or gross morphology

( Robinow and Caten, 1969; Morris, 1976). Mutants were classified into three

groups based on nuclear morphology and distribution and the ability to form

spindles. Three broad classes of mutants displaying defects in the following

7 areas were seen: 1) defective mitoses 2) lack of septa 3) abnormal nuclear

distribution. Cells identified as mitotic mutants were further classified based on

their mitotic indices. Cells unable to enter mitosis at restrictive temperatures were termed nim (never in mitosis) mutants. Cells blocked in mitosis were termed bim (blocked in mitosis) mutants. Cells defective in nuclear migration were termed nud (nuclear distribution) mutants (Morris, 1976). These nine bim mutants in six genes and twenty six mutants nim mutants in twenty three genes provided the basis for future studies into cell cycle regulation in A. nidulans

(Osmani and Mirabito, 2004) (Table 1.1).

Additionally, Aspergillus responds well to traditional arrest treatments including hydroxyurea and anti-tubulin drugs including nocodazole and benomyl allowing for easy arrest and synchronization of viable cultures (Bergen and

Morris 1983). Nuclei can be visualized using DAPI and immunofluorescence and biochemical protocols have been developed. Vector based expression systems exist for exogenous protein expression in the organisms. An alcA based system allows for inducible expression of proteins following integration of transforming

DNA into the genome. Germination of transformed strains on either glucose or ethanol containing media results in transcription repression or induction respectively (Waring et al., 1989; Doonan et al., 1991). An autonomously replicating plasmid based on the AMA1 sequence allows for complementation of recessive mutations and recovery of the complementing DNA (Aleksenko et al.,

1996; Aleksenko and Clutterbuck, 1997). The recent completion of the A. nidulans sequencing project to 13X coverage coupled with the continued

8 development of molecular techniques and construction of cell cycle specific fluorescent protein fusions (H1-GFP, tubulin-GFP, and NPC component-GFP) continue to make A. nidulans an attractive model system for cell cycle research.

1.3. The Cell Cycle

1.3.1. Overview

The maintenance and transmission of a full complement of genetic information in actively dividing cells is essential for the survival of individual cells and the organism as a whole. The growth and division program these cells follow is termed the cell cycle. Our understanding of this essential process has advanced dramatically over the past several decades from a recognition of some of the cytological hallmarks of mitosis to the dissection of complex cell cycle regulatory mechanisms and an ability to mathematically model the cell cycle

(Sveiczer et al., 2004). The scope of this introduction will be limited to a general description of the organization of the cell cycle followed by a discussion of some of the central regulatory mechanisms which govern cell cycle progression including a historical perspective on the role of the cyclin dependant kinases and checkpoint systems.

The eukaryotic cell cycle is composed of two basic phases: preparation for cellular division referred to as interphase and active cellular division, or M phase, which consists of mitosis followed by cytokinesis (Figure 1.5). In most eukaryotic cell cycles, interphase occupies by far the majority of the cell cycle, with active

9 cellular division occupying only a fraction of the cell cycle. In mammalian cells

with a cycle time of twenty-four hours, mitosis would account for approximately

one hour of the total cycle time. Following the separation of , the

cell undergoes cytokinesis, the physical partitioning of the cytoplasm, which

produces two identical cells.

1.3.2. Interphase

Interphase is comprised of three cell cycle phases which includes two gap

phases (G1 and G2) and a DNA replication phase, termed S phase (Figure 1.5).

Movement through interphase occurs sequentially beginning with G1 followed by

S, and finishing in G2. During the two gap phases of the cell cycle the cell

monitors environmental conditions and seeks to attain a size large enough to

undergo division. The cell must be able to maintain a complete complement of

organelles, chromosomes, and other factors required for survival while providing

its daughter cell with enough of these same factors to ensure its survival as well.

If the cell attains appropriate size and environmental conditions are favorable, a

cell will commit itself to S phase at the end of G1. Failure to meet these requirements results in the cell exiting cell division and entering a state termed

G0. The cell may remain suspended in G0 for extended periods of time until

conditions are favorable for reinitiating the cell division cycle.

The central role of the cell cycle is to ensure the faithful transmission of a

complete set of genetic information to both cells produced following cytokinesis.

During S phase a cell must replicate all chromosomes a single time and ensure

10 that the chromosomes are undamaged. The presence of damaged DNA results

in an inhibition of mitotic entry until all DNA can be repaired (Schultz et al., 2000;

Nyberg et al., 2002; Cuddihy and O'Connell, 2003). The cell must also avoid re-

replication of chromosomes prior to mitotic entry. To this end, each S phase is

followed by M and re-initiation of replication is blocked in S phase

and G2 (Heichman and Roberts, 1994; Nurse, 1994). Initiation of DNA replication is controlled primarily by two multiprotein complexes termed the Origin

Recognition Complex (ORC) and the Minichromosome Maintenance complex

(MCM) under the control of CDKs (Cyclin Dependant Kinase) along with additional factors (Nishitani and Lygerou, 2002). These complexes comprise the pre-replicative complexes which “license’ DNA replication origins to allow replication. Following replication initiation in S phases these complexes are converted to post-replicative complexes which make the DNA refractory for re- replication. This block is maintained through mitosis through the action of mitotic

CDKs (Nishitani and Lygerou, 2002). Following the destruction of mitotic cyclins and inactivation of mitotic CDKs pre-replication complexes are reestablished allowing for another round of replication (Nishitani and Lygerou, 2002).

1.3.3. M Phase

M phase consists of two phases of cellular division: mitosis and cytokinesis. The principle focus of mitosis, or nuclear division, is the equal segregation of chromosomes to daughter nuclei. This process involves some of the most striking cytological occurrences the cell undergoes including

11 chromosome condensation, nuclear envelope breakdown and spindle formation.

These processes provide the basis for the further subdivision of mitosis based on

light and electron microscopy studies. These studies have identified five distinct

phases of mitosis. The condensation of chromosomes characterizes the initial

stage of mitosis, called prophase. Each chromosome consists of two sister

chromatids produced during DNA replication in S phase and bound to each other

by a multi-protein complex termed cohesin. At the onset of the next phase,

prometaphase, the nuclear envelope breaks down. Eukaryotic cells which do not

undergo nuclear envelope breakdown, A. nidulans included, are classified as undergoing a closed mitosis. Some of the regulatory consequences of this difference will be discussed later in this thesis. The loss of separation between nucleus and cytoplasm allows the kinetochore microtubules of the mitotic spindle, previously in the cytoplasm, to begin interacting with the chromosomes. During metaphase, chromosomes align along the metaphase plate to allow for equal segregation of chromosomes to both of the poles. To ensure equal chromosome segregation, each chromosome must make a bipolar attachment to spindle microtubules emanating from the spindle poles. A dramatic loss of sister chromatid cohesion signals the beginning of anaphase A. The chromatids are moved toward the spindle pole bodies at opposite ends of the cell. In the second half of anaphase, termed anaphase B, the poles separate with an elongation and sliding of the spindle microtubules. During telophase, the separated sister chromatids arrive at the poles and the nuclear envelope begins to reform around the chromosomes located at both of the poles. Completion of telophase

12 produces a large bi-nucleated cell. The final step of M phase, cytokinesis,

resolves the cell into two identical cells through the process of cytoplasmic

cleavage.

1.3.4. Regulation

1.3.4.1. 2001 Nobel Prize: The awarding of the 2001 Nobel Prize for

Physiology or Medicine jointly to Leland Hartwell, Paul Nurse and Timothy Hunt

“for their discoveries of key regulators of the cell cycle” recognized the significant advancement made in our understanding of the cell cycle over the last several decades. Much of our current understanding of the cell cycle is based on 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 studies included: the identification of cyclin in starfish oocytes by Hunt and colleagues (Evans et al., 1983), roughly parallel genetic screens conducted in budding yeast (Hartwell et al., 1974), fission yeast (Nurse et al., 1976), and A. nidulans (Morris, 1976), and the identification and purification of maturation promoting factor (CDK1 / cyclin B) from frog oocytes

(Masui and Markert, 1971; Gautier et al., 1988; Lohka et al., 1988). For the purposes of this work, specific emphasis will be placed on mechanisms that regulate mitosis.

1.3.4.2. Checkpoints: A core feature of the cell cycle is the dependence of late events on the proper completion of earlier events. For example, entering

13 mitosis prior to successful completion of S phase would produce disastrous

consequences for the cell, as each cell would be unable to receive a full

complement of chromosomes. In a review of cell cycle controls in 1989 Leland

Hartwell and Ted Weinert examined the two models put forward to explain this

dependence (Hartwell and Weinert, 1989). The first model viewed the cell cycle

as a biochemical system in which substrates produced in early stages were

required for completion of later events. In the second model, a control system

was postulated to monitor the cell cycle for successful completion of earlier

events before allowing cells to proceed. Hartwell and Weinert argued for the second model and identified several critical transitions where they said this control mechanism could exert influence (Hartwell and Weinert, 1989). Hartwell and Weinert argued that the ability to identify conditional mutants, chemicals, and other conditions which allowed cells to bypass normal progression represented a

“relief of dependence” which supported the presence of inhibitory control mechanisms which the authors termed “checkpoints” (Hartwell and Weinert,

1989). Today checkpoints are universally accepted as the primary controls governing cell cycle progression.

Three major sites of potential cell cycle arrest exist within the eukaryotic

cell cycle. First, a cell will commit to S phase only if it has achieved an

appropriate size and if environmental conditions favor cellular division. Failure to

meet these requirements results in the cell entering a nonproliferative state called

G0 until conditions again favor cell growth and division. Second, the DNA

Damage and DNA Replication Checkpoints prevent cells from entering mitosis in

14 the presence of unreplicated or damaged DNA. If either of these is present a cell will remain in interphase until DNA replication and repair is complete. Third, during mitosis, a cell will not progress into anaphase if the chromosomes have failed to align properly on the mitotic spindle. This arrest, mediated by the

Spindle Checkpoint, prevents an unequal distribution of chromosomes between parent and daughter cell at cytokinesis. Due to the fact that the ability to arrest cells in metaphase, through activation of the Spindle Checkpoint, is utilized in a set of experiments described in this thesis I will provide a brief overview of the regulatory machinery governing this checkpoint.

Following DNA replication, sister chromatids are linked by a protein complex termed cohesin (Michaelis et al., 1997; Uhlmann and Nasmyth, 1998;

Tanaka et al., 1999; Toth et al., 1999; Ciosk et al., 2000; Tanaka et al., 2000;

Haering et al., 2002). Sister chromatid cohesion persists until anaphase, at which point the cohesin subunit Scc1 is cleaved by a proteolytic termed

Separase, resulting in a loss of chromatid cohesion (Zou et al., 2002) (Ciosk et al., 1998; Uhlmann et al., 1999; Uhlmann et al., 2000; Zou et al., 2002; Rao et al.,

2001; Tomonaga et al., 2000) Separase is maintained in an inactive state, bound to Securin, until all chromosomes have aligned properly on the mitotic spindle (Zou et al., 1999; Stemmann et al., 2001). Once this criterion is met, the

APC (Anaphase Promoting Complex), an E3 ubiquitin , targets Securin for proteolysis (Yu et al., 1998; Yu, 2002). Loss of Securin releases Separase to cleave cohesin and permits anaphase progression.

15 The spindle checkpoint blocks activation of the APC until all chromosomes

have made bi-polar spindle attachments through inhibition of CDC20, thought to

be a targeting subunit of the APC. Inhibition of CDC20 is accomplished by a

group of checkpoint proteins termed the mitotic checkpoint complex, which is

comprised of Mad2, BubR1, Bub3, and Cdc20 (Bharadwaj and Yu, 2004).

Proper alignment of all chromosomes inactivates inhibitory signals which had

been originating from unoccupied kinetochores and dismantles the mitotic

checkpoint complex, resulting in APC activation and anaphase onset (Bharadwaj

and Yu, 2004).

1.3.4.3. Identification of cyclin dependant kinases: While studying protein expression in sea urchin oocytes, Hunt and colleagues identified a protein whose levels rose and fell through the cell cycle (Evans et al., 1983). Early embryonic cell cycles lack the gap phases present in adult organisms, and are characterized by rapid rounds of DNA replication preceded immediately by mitosis. They noted that levels of the protein rose through the division cycle but disappeared just before each cleavage division (Evans et al., 1983). The protein was termed cyclin due to its cyclic nature and Hunt speculated that the expression pattern of the protein may be associated with cell cycle oscillations.

In 1986 experiments performed by Joan Ruderman cemented a central role for

Cyclin in cell cycle progression. Experiments involving the injection of recombinant cyclin A from clam embryos into Xenopus oocytes showed that cyclin was capable of driving these oocytes into meiosis (Swenson et al., 1986).

16 These experiments also indicated the degree to which the core cell cycle

machinery was conserved between organisms.

Work on cyclin was occurring roughly in parallel with work to characterize

the mysterious Maturation Promoting Factor (MPF), first identified in oocytes

from the frog Rana pipiens (Masui and Markert, 1971). By 1967 Masui had

demonstrated that addition of progesterone to frog eggs induced entry into

meiosis (Masui, 1967). Using these activated eggs Masui and colleagues

demonstrated that injection of cytoplasm from progesterone activated eggs into

inactivated oocytes produced rapid entry of the recipient oocyte into meiosis

(Masui and Markert, 1971). Based on this finding they concluded that the

cytoplasm of activated eggs contained a factor capable of promoting maturation,

and the attempts to purify and identify the components of MPF began. In 1988

MPF was purified from frog oocytes and determined to consist of two subunits of

32 kD and 45 kD in size (Lohka et al., 1988). MPF purified from starfish ( Labbe

et al., 1989a; Labbe et al., 1989b) and clams (Draetta et al., 1989) contained

subunits of the same sizes as those in frogs.

Two temperature sensitive mutant screens conducted by Leland Hartwell

in S. cerevisiae and Paul Nurse in S. pombe identified an array of cdc (cell

division cycle) mutants, which exhibited defects in cell cycle progression at restrictive temperatures (Hartwell et al., 1974; Nurse et al., 1976). In the screen conducted by Nurse, a mutation termed cdc2 was identified. Nurse and colleagues demonstrated that Cdc2 was required for initiation of both S and M phases (Nurse et al., 1976; Nurse and Bisset, 1981). Complementation of the ts-

17 phenotype using both S. pombe and S. cerevisiae DNA libraries resulted in the cloning of Cdc2 and the identification of the cdc28 gene previously identified by

Hartwell as its homologue (Beach et al., 1982). Continuing work would demonstrate that Cdc2 and CDC28 were protein kinases and that their activity was maximal at M (Simanis and Nurse, 1986). Cloning of a human Cdc2 cDNA by complementation of the S. pombe cdc2 mutation further emphasized the remarkable conservation of this core cell cycle regulator (Lee and Nurse, 1987).

A Cdc2 specific antibody was used to identify the 32 kD protein in MPF as the human homologue of Cdc2 (Gautier et al., 1988). In addition to isolating Cdc2, the screen also isolated its activating cyclin partner (Booher and Beach, 1987;

Booher and Beach, 1988; Moreno et al., 1989). Work from Paul Nurse’s lab also identified the core regulators of CDK1 activation during mitotic entry including the mitotic inhibitory kinase and the Cdc2 activating Cdc25 phosphatase

(Nurse and Thuriaux, 1980; Russell and Nurse, 1986; Russell and Nurse, 1987;

Gould and Nurse, 1989; Moreno et al., 1990). At the 1991 Cold Spring Harbor

Symposium on the cell cycle it was determined that kinases which associated with cyclins would be termed CDKs (Cyclin Dependant Kinases). Cdc2, CDC28, and human Cdc2 are now referred to as CDK1.

Work over the following years further cemented the central role of CDKs in driving cell cycle progression. Work in yeast has demonstrated that a single

CDK, CDK1, drives major cell cycle transitions through its interaction with nine different cyclins. In higher eukaryotes four different CDKs (CDK1, CDK2, CDK3,

CDK4) associate with phase specific cyclins to drive cell cycle events with CDK1

18 forming a complex with cyclin B to drive mitotic entry. 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 et al., 2003),

and the burgeoning family of NIMA-related kinases (O'Connell et al., 2003).

This presentation of a historical perspective on the isolation and

characterization of the central cell cycle regulator CDK1 serves to illustrate

several points. First, the cell cycle regulatory machinery is highly conserved.

While this introduction highlights only CDK1, the vast majority of core regulators

of the cell cycle exist in fungi through to humans. Second, the use of simplified

model systems and organisms to dissect complex regulatory pathways can be

highly effective. Third, the use of genetics has been central to developing our current understanding of cell cycle regulatory mechanisms. The degree of conservation of the cell cycle machinery supports a continued central role for genetics and model genetic organisms in the study of the eukaryotic cell cycle.

1.4. NIMA protein kinase

The importance of the CDK has been well established in eukaryotic systems ranging from fission yeast to humans. However, a second class of cell cycle regulated protein kinase has been isolated in A. nidulans. The NIMA protein kinase is distinct from CDK1 and related CDKs in that it does not require

19 interaction with cyclins in order to be activated, however, it has been shown to be required, along with active CDK1, for entry into M in A. nidulans.

1.4.1. Isolation

In a screen for temperature sensitive mutants of A. nidulans, N. Ronald

Morris identified a class of mutants, termed nim mutants, which when grown at restrictive temperature, failed to enter mitosis (Morris, 1976) (Figure 1.6). These mutants lacked the ability to form spindles or undergo chromosome condensation

(Morris, 1976). nimA, originally isolated as four different alleles, nimA1, nimA5, nimA7, and nimA9, encodes a serine/threonine specific protein kinase which is required for mitotic entry in A. nidulans (Figure 1.6). While incubation of the temperature sensitive nimA5 allele blocks mitotic entry, shifting this mutant to permissive temperature results in immediate mitotic entry (Oakley and Morris,

1983). Additionally, the study of NIMA related kinases in higher eukaryotes has identified a range of kinases, some with mitotic specific functions.

1.4.2. Functional regions within NIMA

nimA encodes the NIMA protein kinase, a serine/threonine specific protein kinase 699 amino acids in length with a molecular weight of 79 kD (Osmani et al.,

1988b; Lu et al., 1993). The kinase consists of two functional domains; an amino-terminal catalytic domain and a carboxy-terminal non-catalytic domain

(Figure 1.7).

20 The catalytic domain is necessary for NIMA kinase function. Kinase negative versions of NIMA have been produced in two ways. First, truncations within the NIMA kinase domain extinguish activity. Second, the point mutation of single essential residues within the kinase region also extinguishes activity.

NIMA contains two residues within the catalytic domain known to be essential for a broad range of kinases (Steinberg et al., 1993). Mutation of either of these residues, a threonine at position 199 or a lysine at position 40 produce an inactive kinase; (Taylor et al., 1990; Lu et al., 1993; Pu et al., 1995). This ability to produce kinase inactive forms of NIMA has been utilized in a Two-hybrid screen to identify NIMA interacting proteins (Osmani et al., 2003).

Overexpression of NIMA forces cells into premature mitosis characterized by condensed DNA and establishment of mitotic spindles (Osmani et al., 1987).

However, expression of the catalytic domain on its own has no effect on cell cycle progression (Pu and Osmani, 1995). This result highlights the importance of the non-catalytic domain of NIMA. While the amino-terminus encodes a functional kinase domain, the carboxy-terminus is critical for regulating this catalytic function. Several functional motifs within this region highlight these overlapping regulatory systems. First, the timely proteolytic degradation of NIMA is essential for normal cell cycle progression (Pu and Osmani, 1995). 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, and Nurse 1994; Osmani and Pu 1995). Sequences rich in proline, glutamic acid, serine, and threonine (PEST) have been identified in this region (Figure 1.7).

21 PEST sequences are believed to be involved in facilitating rapid degradation of

the proteins in which they are contained (Rogers et al., 1986; Yaglom et al., 1995;

Rechsteiner and Rogers, 1996; Roth and Davis, 2000). Expression of NIMA

which lack these PEST sequences produces a version of NIMA which is far more

stable than normal NIMA (O'Connell et al., 1994; Pu and Osmani, 1995).

Expression of these stable forms of NIMA is highly toxic and is capable of

inducing chromosome condensation in A. nidulans and higher

eukaryotes(O'Connell et al., 1994; Pu and Osmani, 1995). Additionally,

expression of these forms of NIMA blocks mitotic exit in A. nidulans (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 (Figure 1.7). Intriguingly,

like nimA5 and nimA7, the nimA1 temperature sensitive allele blocks cells in G2

at restrictive temperature (Pu et al., 1995; Wu et al., 1998) Unlike, these other alleles, the nimA1 mutations is in the carboxy-terminus and possesses a basal level of activity at restrictive temperature (Pu et al., 1995). The nimA1 mutation

disrupts one of the nuclear localization signals (Pu et al., 1995), and NIMA1 is

cytoplasmic at restrictive temperature but will enter the nucleus in the presence

of the mutant versions of nucleoporins SONA and SONB (Wu et al., 1998; De

Souza et al., 2003). These results suggest a role for these nuclear localization

signals in regulating NIMA function.

22 Finally, two coiled coil domains exist just outside of the catalytic domain

(Figure 1.7). 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 which are 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 et al., 1995; Chiu et al., 2004; Nikolay et al., 2004).

The coiled coil regions in NIMA represent potential proteins interaction sites.

These regions may play a role in the interaction of NIMA with other cell cycle regulatory proteins or a potential NIMA activating kinase. It has been demonstrated that overexpression of the non-catalytic terminus of NIMA produces the same phenotype as overexpression of kinase inactive NIMA. In both cases, A. nidulans transformants arrest in late G2 (Lu and Means, 1994).

This dominant-negative phenotype is believed to be a result of competitive

inhibition. In other words, mutant NIMA constructs out compete endogenous

functional NIMA kinase for substrates essential for mitotic initiation. These

results suggest that the carboxy-terminus of NIMA, potentially the coiled coil

regions, is the site of protein-protein interaction.

23 1.4.3. NIMA fluctuations through the cell cycle

NIMA protein levels and corresponding kinase activity are tightly cell cycle

regulated. The precise regulation of NIMA activation is critical considering

NIMA’s potent mitosis inducing potential. Expression of NIMA can induce

disastrous premature mitoses (Osmani et al., 1988b). NIMA protein levels

remain low throughout S phase (Ye et al., 1995). NIMA levels rise through G2 to reach a maximum at mitotic entry before decreasing rapidly through mitosis (Ye et al., 1995) (Figure 1.8). The level of NIMA mRNA also reaches a maximal level at mitosis (Osmani et al., 1987) (Figure 1.8). The mechanisms underlying these changes in mRNA and NIMA protein levels are not well defined. For example, either increased translation rates or decreased protein degradation could account for increased NIMA levels at mitosis (Osmani and Ye, 1996).

The exact mechanisms underlying NIMA degradation are also unknown.

During interphase, phosphorylated, but not fully activated forms of NIMA accumulate. These forms continue to accumulate in the absence of active CDK1

(Osmani et al., 1991a). Upon entry into mitosis, fully activated NIMA is targeted by aggressive proteolysis (Pu and Osmani, 1995; Ye et al., 1995). The APC represents a good candidate for a regulator of NIMA proteins levels. 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 (Ye et al., 1996). It follows that the APC should also target the

NIMA kinase for destruction to allow for mitotic exit. This idea is supported by the fact that mutations in two different APC components, bimE and bimA, have

24 been shown to be able to override a nimA5 G2 arrest to allow some mitotic

events (Osmani et al., 1988a; Ye et al., 1998)in a NIMA dependant manner.

Considering that APC mutants were unable to overcome a complete loss of nimA

function (Ye et al., 1998), their ability to partially override a nimA5 induced arrest occurs presumably by allowing partially functional NIMA to accumulate to a level high enough to induce some mitotic events. It has been proposed that the hyperphosphorylation of NIMA at the G2 to M transition may act as a ubiquitin-like signal for proteolysis, the multiple phosphorylations effectively "signaling" appropriate proteolytic machinery, probably the APC, to trigger NIMA degradation (Ye 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 (O'Connell et al., 1994; Pu and

Osmani, 1995).

1.4.4. Phosphorylation and NIMA activation

In addition to the modes of regulation described above, NIMA kinase

activity is regulated at the level of phosphorylation. During interphase NIMA is

initially unphosphorylated and inactive as a kinase. Later in G2 NIMA is phosphorylated and displays a basal level of activity toward β-casein (Ye et al.,

1995). This phosphorylation event probably represents phosphorylation of threonine 199, a residue required for NIMA activity, either by autophosphorylation or by the action of a NIMA activating kinase (Pu et al., 1995; Ye et al., 1995). At the G2-M transition NIMA is hyperphosphorylated and becomes MPM-2 reactive,

25 in a CDK1 dependant manner (Ye et al., 1995). It is this hyperphosphorylated

form of NIMA which is present during mitosis, and which is rapidly degraded.

This fact confines maximal NIMA activity to a defined period of early mitosis.

1.4.5. Active NIMA is required for the G2-M transition

Active NIMA kinase is required for all cytological aspects of mitosis in A. nidulans; (Oakley and Morris, 1983; Bergen et al., 1984; Osmani et al., 1987).

Early experiments investigating the role that nimA played in cell cycle progression made use of expression vectors containing the inducible alcA promoter. These expression studies demonstrated that exogenous NIMA expression could drive A. nidulans cells into mitosis from any stage of interphase

(Osmani et al., 1988b). This mitotic arrest is characterized by chromatin

condensation and mitotic spindle formation and elongation (Osmani et al., 1988b).

The requirement of NIMA function for mitotic onset was clearly illustrated in

experiments utilizing the temperature sensitive nimA5 allele. Inactivation of

NIMA5 by incubation at restrictive temperature resulted in accumulation of cells

at G2. The essential cell cycle role for the NIMA kinase is most clear in studies

involving the use of Aspergillus strains possessing the conditional nimA5 mutation. The nimA5 mutation is a temperature sensitive mutation which results in inactivation of NIMA at the restrictive temperature. These studies illustrate that

mutant nimA5 Aspergillus strains with fully activated p34cdc2/cyclin B are unable

to progress past G2 when shifted to restrictive temperature (Osmani et al., 1991a).

Surprisingly, these cells contain fully active CDK1 which is bound to cyclin B and

26 is active as a H1 kinase (Osmani et al., 1991a; Pu et al., 1995). More recent

studies have suggested that NIMA regulates CDK1 not at the level of activity but

by restricting the availability of substrates through regulation of the subcellular

localization of CDK1 (Wu et al., 1998; De Souza et al., 2003).

1.4.6. NIMA interacting proteins and targets

During the last few years several NIMA interacting proteins have been

identified. These NIMA interacting proteins highlight multiple roles for the NIMA kinase in chromatin condensation, regulation of microtubule dynamics, and alteration of nuclear pore complex properties at mitosis.

A hallmark of NIMA is its ability to induce chromosome condensation in A.

nidulans as well as mammalian cells (Osmani et al., 1988b; Lu and Hunter,

1995b; Ye et al., 1995) The identification of histone H3 reveals a potential

mechanism by which NIMA could induce chromosome condensation.

Phosphorylation of histone H3 has been identified as being important for

chromatin condensation (Guo et al., 1995; Hendzel et al., 1997; Van Hooser et

al., 1998; Sauve et al., 1999; Wei et al., 1999). In Tetrahymena, phosphorylation

of Serine 10 is required for chromosome condensation (Wei et al., 1999). In A.

nidulans, NIMA activity is required for Serine 10 phosphorylation (De Souza et al.,

2000). Moreover, NIMA expression induces H3 phosphorylation in S phase

arrested cells and NIMA can phosphorylate serine 10 of H3 in vitro (De Souza et

al., 2000).

27 A screen for mutants capable of suppressing the nimA1 mutation identified

two nucleoporins, SONAGLE2/RAE1 and SONBNUP98 (Wu et al., 1998; De Souza et al., 2003). The interaction of NIMA with nucleoporins suggests a role for NIMA in remodeling the nuclear pore complex to allow mitotic specific regulators, including CDK1 to gain access to the nucleus during a closed mitosis (Wu et al.,

1998; De Souza et al., 2003).

Finally, a yeast two hybrid screen utilizing different forms of NIMA as bait

identified TINA as a strong NIMA interacting protein (Osmani et al., 2003). TINA is a SPB associated protein which interacts with NIMA during G2 and potentially regulates the formation of astral microtubules and interaction between adjoining spindles TINA (Osmani et al., 2003).

1.4.7. NIMA related kinases and cell cycle control

The identification of nimA and the characterization of its essential mitotic specific function in A. nidulans initiated a search for NIMA homologues in other

organisms. Experiments which demonstrated that NIMA overexpression in

human or yeast cells could induce chromosome condensation (O'Connell et al.,

1994; Lu and Hunter, 1995a), and that expression of kinase negative forms of

NIMA could delay mitotic entry in human cells (Lu and Hunter, 1995a) supported

the idea that a NIMA-like regulatory system existed in other organism. To date

the only true functional homolog of NIMA capable of complementing a nimA

(nimA5) mutation in A. nidulans is nim-1 from N. crassa (Pu et al., 1995). Nim-1

was isolated by a low stringency hybridization screening of a N. crassa genomic

28 library. Hybridizing fragments were isolated and were tested for their ability to

complement nimA5 (Pu et al., 1995).

The sequencing of budding and fission yeast and human genomes has resulted in the identification of a family of NIMA-related proteins in other eukaryotes based primarily on their homology to the NIMA kinase domain

(O'Connell et al., 2003) (Figure 1.9). To date, a single NIMA-related kinase has been identified in fission and budding yeast (Fin1p and KIN3 respectively), while eleven different NIMA related kinases (Neks) have been identified in humans

(O'Connell et al., 2003) (Figure 1.9). As noted previously, the carboxy terminus of NIMA is characterized by the presence of coiled-coil and PEST domains.

Many of the NIMA-related kinases contain similar domains in their carboxy- termini, while some homologues contain additional protein domains (Figure 1.9).

The importance of NIMA function at 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.

Nek2 is a centrosome associated kinase which exists in two isoforms (Uto et al., 1999). Nek2 function is important for centrosome function and establishing a bipolar spindle. Specifically, overexpression of Nek2 results in centrosome splitting (Fry et al., 1998b) C-Nap1has been identified as a Nek2 target with a role in centrosome cohesion and establishing a mitotic spindle (Fry et al., 1998a;

Mayor et al., 2000; Mayor et al., 2002; Faragher and Fry, 2003) . An additional

Nek2 target, Hec1, binds to kinetochores and is involved in the spindle

29 checkpoint and regulating kinetochore attachments through its interactions with

SMC proteins and centromere protein Ctf19p (Zheng et al., 1999; Chen et al.,

2002; Martin-Lluesma et al., 2002; DeLuca et al., 2003; Hori et al., 2003;). Kin3, the S. cerevisiae NIMA related kinase has also been shown to interact with Hec1

(Chen et al., 2002). While cells lacking Kin3 are able to survive, the expression of a version of Kin3 equivalent to NIMA7 results in severe defects in chromosome segregation (Chen et al., 2002). Finally a recent study has identified a direct interaction between the spindle checkpoint component MAD1 and Nek2 (Lou et al., 2004).

In addition to Nek2 two additional Neks, Nek6 and Nek9, have potential mitotic specific regulatory roles (Kandli et al., 2000; Hashimoto et al., 2002;

Holland et al., 2002; Roig et al., 2002; Yin et al., 2003). Nek 6 is phosphorylated and activated at mitosis (Yin et al., 2003). Expression of kinase negative forms of Nek6 results in a metaphase arrest followed by apoptosis (Yin et al., 2003).

Nek 9 is also phosphorylated and activated at mitosis (Roig 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 et al., 2002). Interference with

Nek9 function by antibody injection produces chromosome segregation defects

(Roig et al., 2002). Finally, Nek 9 binds to and activates Nek6 (Roig et al., 2002;

Yin et al., 2003).

A NIMA homologue termed Fin1p has been characterized in S. pombe

(Krien et al., 1998). Like NIMA, Fin1p is capable of inducing chromatin

30 condensation in the absence of CDK1 activity and undergoes cell cycle mediated

fluctuations (Krien et al., 1998), however, Fin1p levels appear to peak at the

metaphase to anaphase transition, a point later in mitosis than NIMA (Krien et al.,

2002). Additionally, while not lethal, loss of fin1 does produce severe nuclear envelope perturbations and these cells display synthetic lethality with mutants of the mitotic spindle checkpoint (Krien et al., 1998; Krien et al., 2002). Additional

ts- fin1 mutants displayed defects in spindle pole body microtubule nucleation and failure to recruit 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 suppresses the activity of SIN (Septation Initiation Network) components at “old” SPBs (Grallert et al.,

2004)

While, no true NIMA functional homologue has been identified outside of filamentous fungi, a number of the NIMA-related kinases do share a common theme of regulation. Specifically, the NIMA family of related kinases appears to generally regulate establishment and maintenance of the mitotic spindle apparatus.

1.5. Aims

The overall goal of this work was to characterize the roles which potential

NIMA interacting proteins may play in governing cell cycle events in A. nidulans.

The primary focuses of this thesis are tinC and tinD, two of the potential NIMA interacting protein genes isolated in a yeast Two-hybrid screen.

31 The specific aims of the studies described in this thesis were several fold:

i) to reconfirm and better characterize the nuclear division defect observed for

tinC and tinD, ii) to determine whether expression of full length tinC and tinD produce cell cycle specific defects, iii) to determine the subcellular localization of

TINC and TIND, iv) to determine whether TINC and TIND interact with NIMA in A. nidulans, and v) to determine the functions of TINC and TIND in A. nidulans.

32

(Reproduced from Nature Reviews Genetics, Vol. 3, Casselton, L. and Zolan, M., The art and design of genetic screens: filamentous fungi, pp. 683-697, Copyright 2002, with permission from the author.)

Figure 1.1. 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.

33 Conidiospore

A C Conidiospore

Secondary sterigmata Primary sterigmata

Vesicle B

Foot cell

Germling

Figure 1.2. Asexual life cycle 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. The mature asexual reproductive structure, the conidiophore, develops from these cells in a stepwise fashion. An aerial hyphal branch emerges from the foot cell and extends to a certain height, at which time the tip swells to form a vesicle. Primary and secondary sterigmata develop from t vesicle. Mitotic division and cytokinesis at the apex of the secondary sterigmata produces chains of conidia. A. nidulans requires an air interface in order to form conidiophores as A. nidulans germinated in liquid culture fail to produce these structures.

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. The range of different colors, including chartreuse, fawn, white, and yellow, is produced by mutations in genes regulating spore color. These color mutations provide useful tools during genetic manipulations.

35 number ofnuclei.Thelar determine thenumberofmitoseswhich septation occursduringtheseearlyrounds polarized growthwithsynchronousr is amulti-nucleateorganism.During Figure 1.4.Polarizedgrow Mitotic Nuclear Division gest cellpicturedhasundergonef th andnucleardivisionin Polarized Grow ounds ofnucleardivis germination theorganismundergoeshighly 36 hav of nucleardivision,itispossibleto e occurredinacellbas (Image court A. nidulans th i on. Sinceno our mi e s y

Dr. Stephen totic divisions. ed onthe .

A. nidulans A.Osmani)

M G2 M PHASE

G1

INTERPHASE S

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 Wild type nim Mutant

NIMA NIMA

NIMA NIMA

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 mitosis. In the absence of functional NIMA cells

fail to undergo nuclear division and arrest in G2. This failure to enter mitosis was

termed a nim (never in mitosis) phenotype by Ron Morris in a screen he conducted to identify nuclear division, nuclear movement and septation mutants in A. nidulans.

38 Nuclear Localization Signals

Catalytic Domain

Potential coiled coil domains PEST sequences

0 100 200 300 400 500 600 700 (aa)

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

(Adapted from EMBO J., Vol. 14, Ye, et al., The NIMA protein kinase is hyperphosphorylated and activated downstream of p34cdc2/cyclin B: coordination of two mitosis promoting kinases, pp. 986-994, Copyright 1995, with permission from Nature.)

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 maximum at mitosis. During G2 NIMA is phosphorylated and activated as a

kinase. NIMA becomes hyperphosphorylated and fully activated at the G2-M transition. During mitosis NIMA is aggressively proteolyzed, resulting in falling protein levels and kinase activity through mitotic exit.

40

(Reproduced from Trends in Cell Biology, Vol. 13, O’Connell, et al., Never say never. The NIMA-related protein kinases in mitotic control, pp. 221-228, Copyright 2003, with permission from Elsevier.)

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 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 γ 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 ,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 scaffold 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 Fungal Genetics and Biology, Vol. 41, Osmani, S. A. and Mirabito, P. M., The early impact of genetics on our understanding of cell cycle regulation in Aspergillus nidulans, pp. 401-410, Copyright 2004, with permission from Elsevier.)

Table 1.1. List of A. nidulans mitotic 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 (Qiagen or Bio Rad).

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). Minipreps were performed using a miniprep plasmid kit according to the manufacturer’s instructions (Qiagen).

43 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. 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.

44 2.1.4. Primers

Oligonucleotide primers for sequencing, tagging, and cloning were designed using Primer Designer Ver. 2.0 software (Scientific and Educational

Software). Table 2 contains a list of primers used for cloning and PCR based tagging protocols.

2.1.5. Automated fluorescent sequencing

Automated sequencing was performed using either the ABI Prism DNA

Sequencing Kit - Big Dye Terminator Cycle Sequencing Ready Reactions (Perkin

Elmer Applied Biosystems) and the 377 Automated Fluorescent Sequencer

(Perkin Elmer Applied Biosystems) or the Thermo Sequenase dye terminator cycle sequencing pre-mix kit (Amersham Life Science) and the 373A Automated

Fluorescent Sequencer (Perkin Elmer Applied Biosystems). DNA for sequencing was prepared by cesium purification or using Qiagen Maxi-prep and Mini-prep kits. The amount of DNA used for sequencing reactions was 500 ng plasmid

DNA or 1 µg cosmid DNA. 4 pmol of sequencing primer was used in each reaction. PCR was performed using a 9700 Thermal Cycler (Perkin Elmer

Applied Biosystems). PCR cycling conditions for sequencing were 96°C for 30 seconds, 55°C for 15 seconds, and 55°C for 4 minutes (30 cycles) followed by a

4°C soak. Additional sequencing was performed by the Plant Microbial

Genomics Facility at The Ohio State University (Columbus, OH). Sample preparation was performed in accordance with guidelines available at: http://www.biosci.ohio-state.edu/~pmgf/services-dnasequencing.htm. DNA

45 constructs were sequenced fully on both of the complementary strands.

Sequencing Analysis software Ver 3.3 (Perkin Elmer Applied Biosystems) or

DNAStar Seqman software (DNAStar) were used for sequence alignments and contig generation.

2.1.6. Bacterial strains

DH5αF’ competent Escherichia coli were used for all general cloning and plasmid amplification applications. Additional strains of bacteria 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.

20 ng supercoiled plasmid DNA was added directly to the cells and mixed gently.

When transforming plasmid DNA from low-melt agarose, the agarose was warmed at 65°C for 15 minutes and 30 µL TCM (10 mM Trizma Base [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 42°C waterbath 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 37°C and 225 rpm. Transformed cells 46 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 37°C 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 37°C 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 -80°C.

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 auxotrophic 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)

47 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 auxotrophic 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 concentration 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 potassium 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

48 6.8] to 12 mM and sodium thiosulfate to 3.2 mM. 40 mM threonine was included

for alcA:: based protein induction.

2.2.2. A. nidulans Strains

Table 2.2 contains an alphabetical list of all A. nidulans strains used in

these studies.

2.2.3. Preparation of A. nidulans conidia stock suspensions

Haploid A. nidulans strains were inoculated at a concentration of 1X107 spores/ml into 4 ml of MAG media (strains carrying uncomplemented pyrG89 were supplemented with uracil and uridine) containing 0.9% agar at 48°C.

Inoculated media was vortexed to mix the suspension and overlaid onto MAG plates. Inoculated 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 minutes 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.

49 2.2.4. Conidiospore Quantitation

Conidial suspensions of A. nidulans were quantitated to allow for accurate

inoculation densities for germination in liquid media. Conidial suspensions were

quantitated 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/ml to determine the concentration of the original suspension.

2.2.5. 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 37°C for 48 hours and then at room temperature for 3 days. 5 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 300 µL of the suspended spores into three screw top vials containing

baked silica. The silica was vortexed briefly to assist in even distribution of the

spores and returned to ice for 30 minutes. The silica was left at room

temperature for 3 to 4 days with the vial lid loosened to allow for complete drying.

Two vials were placed in a room temperature desiccator and one was placed in a

50 4°C 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.

2.2.6. Strain generation by meiotic crossing

Parental 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. Cleistothecia 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 cleistothecia. Once cleaned, cleistothecia were crushed in 0.2% Tween 20 in a

1.5 ml Eppendorf tube to release conidiospores. Conidiospores were plated on

MAGUU to determine whether the strains had crossed. Individual colonies were

51 selected and tested on a range of minimal media plates lacking various

supplements to identify strains with desired genotypes.

2.2.7. Diploid formation

Conidia from two haploid strains with complementary auxotrophic

requirements and different spore colors were floated on the surface of 3 ml of

dH2O in a “heterokaryon formation tube”. To create a heterokaryon tube, 2 ml of

MAGUU media is allowed to solidify at the bottom of a sterile 10 ml flint glass

tube (Fisher Scientific). The media is then overlaid with 3 ml of dH2O and spores

from both strains are floated on the surface. Strains were germinated overnight

at 30°C to allow a mixed myceliuml mat to form. Sterile tweezers were used to

inoculate small pieces of mixed mycelium onto the surface of selective minimal

media plates to force heterokaryon formation. Spores were harvested from the

surface of heterokaryons in 0.2% Tween 20. Spore suspensions were added at

various dilutions to limiting minimal media and plated. Plates were incubated at

30°C to allow colon formation. Diploids were selected on the basis of spore color

(green).

2.3. General A. nidulans techniques

2.3.1. Small scale protein preparation

Conidia were inoculated at roughly 1x106 conidia/ml into 30 ml of YG or minimal media in a sterile Petri dish. Cultures were incubated overnight at 30°C

52 until just before hyphae began to conidiate at the media air interface. Mycelium

was harvested through Miracloth as described for large scale protein preparation.

Mycelium was frozen in liquid nitrogen and dried overnight in a lyophilizer

(Savant). The next day samples were pulverized with toothpick. The mycelium was 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 mycelium. Samples were boiled for 5

minutes prior to analysis by SDS-PAGE.

2.3.2. Large scale protein preparation

Conidia were inoculated into appropriate liquid media at a concentration of

2x106 conidia/ml. Culture flasks were coated with GelSlick (Cambrex Bio

Science) prior to culture addition to prevent mycelium 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 minute at 7,000 rpm in a clinical swinging bucket centrifuge

(Thermo IEC). Packed cell volumes of between 0.4 to 0.6 ml of mycelium

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 sodium chloride, 65 mg/l sodium azide,

20 mL 0.5 M ethylenediaminetetraacetic acid (EDTA) [pH 8.0], 2.1 g/l sodium

53 fluoride). Excess liquid was pressed out of mycelium 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. Frozen

mycelium was stored at 80°C until protein preparation.

Protein was prepared by grinding frozen mycelium in a prechilled mortar

and pestle in bucket of ice in 2.5 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 aprotanin (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, 71

µl/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 Trizma Base, [pH 7.5], 24 µl/ml 10% Nonidet P-40.) /1 g of

mycelium. Ground myceliuml slurries were aliquotted into 1.5 ml Eppendorf

tubes and centrifuged at 14,000 rpm at 4°C for 10 minutes to pellet myceliuml

debris. The supernatant was transferred to fresh Eppendorf tubes and frozen in

liquid nitrogen. Proteins were stored at -80°C until use. Immediately prior to use

protein samples were rapidly thawed in cold water and centrifuged at 14,000 rpm

at 4°C.

54 2.3.3. Small scale genomic DNA extraction

A small amount of spores were scraped from the surface of a colony using

a sterile toothpick. Spores were placed on the wall of a 1.5 ml Eppendorf tube

and microwaved for 2 minutes. Spores were crushed with a piece of sterile

plastic. 100 µl of Miniprep Lysis Solution (Promega) was added and vortexed to

mix. Contents were microwaved for an additional 2 minutes. 100 µl Miniprep

Neutralization Solution was added and vortexed to mix. Samples were

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 was inoculated into appropriate media and incubated overnight in an air incubator at 32°C at 180 rpm. Inoculums 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 mycelium 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

55 lyophilized overnight. Lyophilized mycelium samples were stored at -80°C until

DNA extraction.

To extract genomic DNA, 20 mg lyophilized mycelium 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 (200 mM Trizma Base [pH 8.5],

250 mM NaCl, 25 mM EDTA, 0.5% SDS), 175 µL phenol, and 75 µl chloroform

were added directly to 40 mg of ground mycelium. 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.

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 32°C 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,

56 2.5 µg/ml ribo, trace elements, 4.92 g/l magnesium sulfate), 80 mg bovine serum albumin (BSA), 20 mg novozyme, 100 mg driselase and 100 µl β-glucoronidase.

The resuspended germlings were transferred to a clean sterile flask and incubated in an air incubator at 32°C 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 potassium chloride, 7.35 g/l calcium chloride, 2.09 g/l MOPS, [pH 6.0]).

Transformation was carried out by combining 4 µg DNA, 100 µl protoplasts, and 50 µl of room temperature Solution 4 (250 g/L PEG 8000, 7.35 g/l calcium chloride, 44.7 g/l potassium chloride, 10 ml 1 M Trizma Base [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 magnesium sulfate, 1

µg/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.

57 2.3.6. Immunofluorescence

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 until 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 (45 mM PIPES, 45 mM HEPES, 10 mM EGTA,

5mM magnesium chloride, [pH 6.9])) for 45 minutes 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 minute intervals following digestion for

1.5 hours. For cells germinated in YG media 2X immunofluorescence digest mix

was used. (Digestion was not required for examination of GFP fusion proteins

GFP-SONBNUP98, TINC-GFP, or TIND-GFP). Coverslips were washed three times in PHEM. Primary antibody incubations were carried out by inversion of

58 coverslips in 150 µl of immunofluorescence buffer (PHEM, 0.01% Tween20, and

3% BSA) plus primary antibodies for one hour at room temperature. Primary antibodies used in these studies included: TAT1 mouse monoclonal anti-tubulin

(1:1,000 dilution), 3F10 rat anti-HA (Roche Biochemicals) (1:800 dilution), and α-

TINC rabbit anti-TINC (1:2,000 dilution). Coverslips were washed three times in

PHEM. Secondary antibody incubations were carried out as described for primary antibodies. Secondary antibodies used in these studies included (anti- mouse, anti-rat, and anti-rabbit AlexaFluor 488 and AlexaFluor 594 (Molecular

Probes, Inc.). Following secondary antibody incubation, coverslips were washed three times in PHEM and mounted in 11 µl of an 80% Citifluor solution with 150 ng/µl DAPI.

2.3.7. Staining of mitochondria

Cells were incubated in media containing 25 nM Mitotracker (Molecular

Probes, Inc.) for 30 minutes. Cells were then processed for immunofluorescence as described previously. Digestion times of 30 minutes were typically adequate to visualize mitochondria.

2.4. Isolation of full-length cDNAs by 5’RACE PCR

The nature of 5’ in-frame fusion between the A. nidulans library cDNAs and the pAD-GAL4 domain means that cDNAs recovered from the yeast Two- hybrid screen are frequently 5’ truncated. To verify identify full-length cDNA

59 clones for tinC and tinD we used 5’ RACE PCR. A full-length An-hetC cDNA was also isolated using 5’ RACE PCR.

To generate an A. nidulans cDNA library, A. nidulans total RNA was

prepared from 2 g lyophilized R153 mycelium using Ultraspec RNA Isolation

System (BIOTECX). Lyophilized mycelium was ground using a ceramic mortar

and pestle which was pretreated in 2 % H2O2 for 1 hour to eliminate RNAse activity. Ground mycelium was combined with 40 ml of Ultraspec RNA reagent and placed on ice for 5 minutes. 8 ml chloroform was added and the mycelium was placed on ice for an additional 5 minutes. Mycelium was centrifuged at

14,000 rpm for 15 minutes to remove debris. RNA was precipitated using 2- isopropanol 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 65° 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

60 Poly-A+ mRNA and cDNA synthesis reagents from the Marathon cDNA

Amplification kit (Clontech)

Full-length tinC and tinD cDNAs were isolated by 5’ RACE PCR using the

Touchdown PCR Method as described in the Marathon cDNA Amplification protocol (Clontech). PCR was performed using an AP1 adaptor ligated cDNA

library, an adapter primer (complementary to the adaptor sequence), and gene

specific primers for tinC (AO79) and tinD (AO11). 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..

2.5. Isolation of Genomic clones for tinC and tinD

A full-length genomic clone for tinC was obtained by performing rounds of

PCR using gene specific primers and a sorted A. nidulans chromosome specific

cosmid library (obtained from the Fungal Genetic Stock Center). A. nidulans

possesses eight chromosomes, all of which were represented in the library. To

identify which chromosomes tinC and tinD reside on, cosmid pools were diluted

in TE to a concentration of 50 ng/µL and combined with all other pools for a

particular chromosome. PCR was performed using tinC specific primers AO22

and AO45 or tinD specific primers AO7 and AO40 (200 nM final concentration), 1

µL of the total chromosomal DNA preparation, 10mM dNTPs, 10X Vent

Polymerase Buffer, and 1 U Vent Polymerase. PCR cycling conditions were as

follows: 94°C 5 minutes, 25 cycles (94°C 45 seconds, 55°C 45 seconds, 72°C

1.5 minutes), 72°C 7 minutes. The presence of tinC or tinD on a particular

61 chromosome was determined by running 4 µl of completed PCR reaction on a

0.8% agarose gel and looking for the presence of a PCR product. The

approximate size of the expected PCR product was determined by performing a

parallel PCR reaction using primers gene specific primers with the tinC or tinD cDNAs recovered from the Two-Hybrid screen as the template. After specific chromosomes were identified for tinC and tinD, tinC and tinD containing cosmids were identified by additional rounds of PCR using chromosome specific cosmid pools. Cosmid DNA was prepared and tinC and tinD was completely sequenced on both strands by primer walking. Gene specific primers with 5’ restriction linkers were used to PCR genomic constructs corresponding to TINC (JDpAL3A and JDpAL3C). ∆N-TINC (JLNIPCA and JDpAL3C), TIND (AO123 and

AO119..), and ∆N-TIND (AO120 and AO119). Note that for ∆N-TINC and ∆N-

TIND constructs, the 5’ primer included a transcriptional start site. PCR setup was as follows: 300ng cosmid DNA, 2.5 U Pfu Polymerase (Stratagene), 1X Pfu

Buffer (Stratagene), 200 nM primers and 200 µM each dNTP. PCR cycling conditions were as follows: 95°C 2 minutes, 30 cycles (95°C 30 seconds, 65°C

30 seconds, 72°C 4 minutes), 72°C 10 minutes. PCR products were cloned as

Kpn1 / BamH1 fragments into either pAL5 (TINC) or pAL3 (TIND).

2.6. TINC antibody

Peptide antibodies were generated against a sixteen amino acid TINC-

specific peptide (KGEYEPQGYERQGSQL). Following synthesis, the peptide

was linked to KLH and rabbits were immunized. Peptide synthesis and

62 production of α-TINC affinity purified anti-serum was performed by Bethyl

Laboratories, Inc., Montgomery, TX.

Strain R153 was germinated in YG media until early log phase, harvested in Stop Buffer, and total protein lysates prepared in HK. 200 µg total protein lysates were subjected to SDS-PAGE and western blotting analysis using α-TINC antibodies.

2.7. HA tagging of tinC and tinD

HA epitope tagged TINC. ∆N-TINC, TIND, and ∆N-TIND constructs were generated using a site directed mutagenesis method developed in our lab ((Wu et al., 1998). The method is based on the Site Directed Mutageneis protocol

(Stratagene). PCR setup was as follows: 100 ng pAL5-TINC or pAL5-TIND constructs (described in previous section), 200µM each dNTP, 1X Pfu Reaction

Buffer (Stratagene), 2.5 U Pfu Turbo Polymerase, and 200 nM each gene specific primers for TINC constructs and TIND constructs (AO135 and AO136).

Note that HA tag was encoded within the primer sequence. PCR conditions were as follows: 95°C 30 seconds and eighteen cycles of 95°C 30 seconds, 65°C 1 minute, and 68°C 20 minutes. PCR Products were digested with Dpn1 enzyme

(Stratagene) for one hour at 37°C and transformed into XL1 Supercompetent

Cells (Stratagene). HA-tagged constructs were isolated and sequenced on both strands to confirm proper HA integration.

63 2.8. GFP tagging of tinC and tinD

TINC was 3’ GFP tagged using a plasmid recombination strategy

described previously (Yu et al., 2000; Chaveroche et al., 2000). A plasmid

containing an 8.8 kb fragment of genomic DNA which included tinC, was

cotransformed into the E. coli strain DY331, following induction of the cells at

42°C for 15 minutes, (Yu et al., 2000) with a 3’GFP-pyrG-zeoR tagging cassette containing 50 bp of tinC sequence immediately upstream of the stop codon and

50 bp of tinC 3’ untranslated region sequence at the 5’ and 3’ends of the cassette respectively. The 3’ GFP tagging cassette was generated by PCR of plasmid which contained a GFP pyrG cassette with primers AO199 and AO200. The recombinant plasmid was recovered and fully sequenced. Plasmid DNA was used for transformation of A. nidulans strain GR5. A. nidulans transformants were selected at random, germinated in minimal media and examined for GFP fluorescence by fluorescence microscopy. Expression of the TINC-GFP fusion protein was confirmed by western blotting.

2.9. alcA driven protein expression in A. nidulans

Genomic DNA fragments corresponding to TINC, ∆N-TINC, TIND and ∆N-

TIND 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.

64 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, 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 40 mM threonine.

2.10. Examination of TINC and TIND localization

For TINC localization, strain JD163 (tinC-GFP) was inoculated into

minimal media glucose at 5x105 condida/ml. 300 µl of inocula was placed on the surface of sterile coverslips and incubated at 30°C to allow germination. Cells were processed for immunofluorescence as described previously. Spindle morphology and DNA condensation were used to monitor cell cycle progression.

To examine endogenous TINC localization, strain R153 was inoculated into YG media at 2x105 conidia / ml. 300 µl of inocula was placed on the surface of

65 sterile coverslips and incubated at 30°C to allow germination. Cells were

processed for immunofluorescence using α-TINC antibodies as described

previously.

For TIND localization strains JD70 (alcA::tinD-HA) or JD164 (tinD-GFP) were inoculated into either minimal media glycerol (for JD70) or minimal media glucose (for JD164) at 5x105 condida/mL. 300 µl of inocula was placed on the surface of sterile coverslips and incubated at 30°C to allow germination. Cells were processed for immunofluorescence as described previously. To visualize mitochondria in cells expressing epitope tagged TIND, cells were costained with

MitoTracker (Molecular Probes) at 25 nM as described previously.

2.11. Deletion of tinC and An-hetC

Several studies have emphasized the utility of bacterial strains containing

inducible recombination machinery to generate recombinant plasmids or cosmids

for the purposes of gene deletion and gene tagging (Yu et al., 2000; Chaveroche

et al., 2000; Swaminathan et al., 2001) We have made use of this strategy to

generate tinC and An-hetC deletion constructs which contain large regions of homologous DNA. The E. coli strains DY380 and DY331 contain temperature

inducible recombination genes exo, bet, and gam (Yu et al., 2000). These

strains display very high recombination efficiencies such that only 50 bp of

homology is sufficient for homologous recombination. Briefly, these E. coli

strains were grown in LB at 30°C to OD600 of 0.4-0.6. The recombination machinery was induced by incubating the cells at 42°C for 15 minutes. Cells

66 were cooled in an ice slurry and spun down and washed in dH2O) twice at 7,000 rpm. Cells were resuspended in 100 µl aliquots and used the same day.

To delete tinC an 8,802 kb genomic fragment containing tinC was cloned into the Litmus 29 plasmid vector as an AflII/AvrII restriction fragment. This plasmid was transformed into induced DY331 bacterial cells (Yu et al., 2000) by electroporation, along with a bifunctional deletion cassette.

The bi-functional deletion cassette was produced by PCR using the pCDA21 plasmid (Chaveroche et al., 2000) and two primers containing 50 bp of homology to regions in the 5’ and 3’ UTR of tinC. PCR was performed using the

Expand Long Template PCR Kit (Roche) according to the manufacturer’s instructions. PCR produced a cassette which contained 50 bp of tinC 5’ UTR homology followed by the zeomycin resistance gene and A. fumigatus pyrG, and ending with an additional 50 bp of sequence homologous to sequence in the tinC

3’ UTR. A recombined vector in which tinC was replaced by the zeoR/pyrG was

recovered. The plasmid was digested with AvrII and AflII to linearize the

construct and transformed into A. nidulans strain GR5. pyrG+ transformants were recovered, streaked to single colonies three times, and genomic DNA obtained. PCR was used to initially confirm which strains were tinC deleted.

Pimer set AO63 and AO74 amplified a 421 bp fragment if endogenos tinC was present. Pimer set AO158 and AO165 amplified a 540 bp fragment if the deletion construct was present. Strains in which a 540 bp fragments was amplified with no 421 bp fragment were regarded as potential tinC deleted strains.

Lack of tinC was confirmed by southern blotting and subjecting 6 M Urea protein

67 lysates from transformants to SDS-PAGE and western blotting with α-TINC

antibodies.

Deletion of An-hetC was achieved in generally the same manner as

described for tinC, with a few exceptions. An An-hetC containing BAC (Bacterial

Artificial Chromosome) was isolated and transformed into uninduced DY380 cells

by electroporation. These cells were then induced for recombination and a

zeoR/pyroA+ cassette was introduced by electroporation. A recombined An-hetC

deleted BAC was recovered. Restriction digest and southern blot analysis

identified Age1 restriction fragment which contained ∆An-hetC. This fragment

was transformed into A. nidulans strain GR5 and pyroA+ transformants slelected.

Genomic DNA was isolated from these transformants and PCR screening was performed using the Expand Long Template PCR Kit (Roche) and primers

AO193 and AO205. Deletion of An-het-C was also confirmed by southern blotting.

Original attempts to generate ∆tinC and ∆An-hetC double deletion strains

involved transforming the An-hetC deletion cassette into strain JD82 and

selecting for pyroA+ transformants, however, no double deleted strains were ever

obtained using this approach. Strains lacking both tinC and An-hetC were

successfully created by crossing ∆tinC and ∆An-het-C strains. I will briefly outline the strategy. Strain JD82 was crossed to SWJ298 to obtain JD116

(pyroA4; pyrG89 + ∆tinC pyr4+; nicA2; chaA1). JD 116 was crossed to JD98 and pyr4+ / pyroA+ progeny were selected. Deletion of tinC and An-hetC in these

68 strains was confirmed by PCR screening and western blotting with α-TINC antibodies.

2.12. ∆tinC and ∆An-hetC phenotype testing

∆tinC strain JD82, ∆An-hetC strain JD98, and ∆tinC/∆An-hetC strains

JD124, JD126, JD127, and JD128 were tested for growth defects along with control strains JD83 and JD93 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 1 M 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 (4 M, 6 M, and 10 M). All growth plates were incubated at 32°C except where indicated.

These strains were also examined for cell cycle defects by DAPI staining and tubulin immunofluorescence. These strains were initially grown in either

YGUU at 32°C. When no defects were observed, strains were germinated at

20°C or in YG containing 1M sucrose or 1M sodium chloride. Additionally, to examine whether loss of tinc and An-hetC function affects nuclear membrane fission, a strain lacking both tinC and An-hetC and which expressed sonA-GFP was created by crossing JD127 to CDS171 to obtain strain JD160. This strain was also examined by immunofluorescence microscopy under the conditions described above.

69 2.13. Crosses between ∆tinC / ∆An-hetC and cell cycle mutants

To determine whether loss of tinc and An-hetC function was synthetically

lethal with mutations in cell cycle regulatory genes, Strains JD124 and JD127

were crossed to a number of strains carrying the following mutations: nimA1,

nimA5, nimA7, nimX1 (CDK1), nimX2, nimX3, nudA5 (cytoplasmic dynein), nudA7, nimT23 (Cdc25), and bimE7 (APC1). These triple 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).

2.14. Examination of nuclear division defects in ∆N-TINC or ∆N-TIND

expressing cells

Strains JD148 (vector control), JD155 (∆N-TINC), JD157 (vector control),

and JD159 (∆N-TIND) were inoculated into minimal media ethanol at 5x105 conidia / ml. 300 µl of innocula was placed on the surface of sterile coverslips and incubated at 30°C to allow germination. Cells were fixed and DAPI stained as described previously.

To quantify the nuclear division defect phenotypes, DAPI staining was

used to classify the cells as having one of four classes of nuclei: 1) single DNA

mass, 2) separated DNA, 3) separated DNA with visible DNA linkage, and 4)

fragmented DNA. 100 cells were characterized for each strain.

70 2.15. Examination of nuclear envelopes and nuclear membrane fission

defects in cells expressing ∆N-TINC or ∆N-TIND

Strains JD157 (vector control), JD158 (∆N-TINC), and JD159 (∆N-TIND)

were inoculated into minimal media ethanol at 5x105 conidia / ml. 300 µl of

inocula was placed on the surface of sterile coverslips and incubated at 30°C to

allow germination. Cells were processed for immunofluorescence as described

previously. Cell cycle stage was determined by spindle morphology, DNA

condensation, and GFP-SONBNUP98 localization.

To quantify the nuclear membrane fission defects for strains JD158 and

JD159, DAPI staining was used to identify cells which appeared to contain separated DNA. GFP-SONBNUP98 localization was then examined to determine

whether the separated DNA masses were contained within a single nuclear

envelope. For strains JD157 and JD159, 100 cells were counted. For strain

JD158, 50 cells were counted.

2.16. Examination of ∆N-TINC and ∆N-TIND localization

For ∆N-TINC localization, strain JD155 was inoculated into minimal media

ethanol at 5x105 conidia / ml. 300 µl of inocula was placed on the surface of

sterile coverslips and incubated at 30°C to allow germination. Cells were

processed for immunofluorescence as described previously using anti-HA

antibodies. Alternately, to examine the accumulation of ∆N-TINC over time, strain CB45 was inoculated into minimal media glucose at 8x105 conidia / ml.

300 µl of inocula was placed on the surface of sterile coverslips and incubated at

71 30°C to allow germination. Coverslips were washed in minimal media without

carbon source and then transferred into minimal media with ethanol. Samples

were removed and process for immunofluorescence using anti-HA antibodies at

1, 3 and 6 hours following the media switch.

For ∆N-TIND localization strain JD73 was inoculated into minimal media glycerol at 5x105 conidia / ml. 300 µl of inocula was placed on the surface of sterile coverslips and incubated at 30°C to allow germination. Cells were processed for immunofluorescence as described previously using anti-HA antibodies.

2.17. TINC and NIMA co immunoprecipitations

For NIMA TINC co-immunoprecipitation experiments, strain R153 was germinated to mid log phase in YG media. Protein was prepared in HK or HK

buffer lacking all phosphatase inhibitors. NIMA was immunoprecipitated from 6

mg of total protein using 30 µL affinity purified α-NIMA antibodies raised against the ANYRED peptide (Ye et al., 1996). Immunoprecipitates were incubated with biotinylated donkey anti-sheep antibodies followed by addition of Streptavidin

MagneSphere Paramagnetic particles (Promega). Following extensive washing in HK and RIPA buffers, immunoprecipitates were subjected to SDS-PAGE and western blotting. Proteins were visualized using either E18 anti-NIMA antibodies or α-TINC antibodies (generated against TINC specific peptide –

KGEYEPQGYERQGSQL). Co-immunoprecipitation experiments conducted from strains carrying the nimA7 mutation were performed in an identical manner with

72 the following exceptions. Strain SO117 was germinated in YG media to early log

phase and shifted to 42°C for 3 hours to inactivate nimA7. The strain was

harvested at this point and total protein lysates created either in HK with all

phosphatase inhibitors (including microcystin) or in HK with all phosphatase

inhibitors except microcystin. Subsequent analysis was performed as described

above.

2.18. Examination of NIMA in ∆N-TIND and ∆N-TIND expressing cells

For NIMA stability experiments, strains D23C, JD161, and JD162 were

inoculated into minimal media yeast lactose plus 40 mM threonine, and grown to

early log phase at 30°C. G2 arrested samples were created by shifting the cultures to 42°C by shaking the culture flasks in a 55°C water bath. Cultures were incubated at 42°C for three hours to allow inactivation of nimT23cdc25 and arrest cells in G2. To generate mitotic samples, cultures were grown to early log phase then shifted to 42°C for three hours. At this point, nocodazole was added to a final concentration of 5 µg/ml (to activate the spindle assembly checkpoint) with continued incubation at 42°C for 10 minutes. Cultures were downshifted to

30°C permissive temperature in an ice bath to allow release of the sample into metaphase. Time course samples were removed at 20, 40, and 60 minutes following downshift to permissive temperature. Protein samples were prepared in HK buffer plus microcystin. NIMA was immunoprecipitated from 5 mg of total protein using 25 µl affinity purified α-NIMA antibodies raised against the

ANYRED peptide. Immunoprecipitates were subjected to SDS-PAGE and

73 western blotting with the E15 NIMA specific antibodies. For western blotting of

NIME and tubulin, 200 µg of sample was subjected to SDS-PAGE and direct western blotting with α-NIME antibodies (E8 IS2) or B512 (Sigma).

2.19. 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.).

Live cell imaging was performed on an inverted microscope (Nikon, Inc.) fitted with a Wallac Ultraview Live Cell Imager spinning disk confocal system (Perkin

Elmer, Inc.). Data collection and analysis on both microscopes was performed using Ultraview image capture software (Perkin Elmer, Inc.).

To examine GFP fusion 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 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). Examination of fixed immunofluorescence samples was performed using a Nikon Eclipse E800 microscope and an UltraPix digital camera (Life Science Resources, Ltd.). Image capture on both microscopes was performed using Ultraview image capture software (Perkin Elmer).

74 2.20. Bioinfomatics 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 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/).

75 Primer Name Primer Sequence 5’ → 3’ ______

AO7 CAC CCG TCT ACG CTT GCC GC

AO11 CTG ACT GCA CAC CCT CGA CG

AO22 ACG CGA GGT CTA GGA ATG AC

AO40 TGT TGT CGG AGC TTC TCC TC

AO45 CGA ATG AGT GGG ATT GCT GC

AO63 ACT CAT CAT GAC GGA CGT GG

AO74 TGC CTA GAC AGT AAG TCC AC

AO79 CAG CTT CAG ACT CTG CGA TTA GTA CC

AO119 TAT GGA TCC CAT TGT GAG AGT TCC TGG TGA GTC

AO120 TTA GGT ACC ATG CTG TCT GCA GCT CAC GGC C

AO123 TTA GGT ACC ATA CTT TCA ACG CAG CAC AAC ACA T

AO135 TAC CCA TAC GAT GTT CCT GAC TAT GCG GGC TAT CCC TAT GAC GTC CCG GAC TAT GCA GGA TAG AGG AGA CCT CGT ATT TGA CTT TGT AG

AO136 TCC TGC ATA GTC CGG GAC GTC ATA GGG ATA GCC CGC ATA GTC AGG AAC ATC GTA TGG GTA GGG CGC CGA ATC TAG AGA AGA GG

AO153 CTG TTA TGC CCA AAG AAA ATC CAG A

AO158 GTG ACC CTG TTC ATC AGC

AO165 AAG CGA GCT GTT GAG ATG GAA G

Continued

Table 2.1. List of primers used for cloning and tagging protocols

76 Table 2.1. continued

AO193 AAG TGA GGC AGA TGG CAA TG

AO199 CGC AAG GCG AAC ACG GCC AGC GTG GTG ATG AGG ACA GGA GTT ACG GGT ACC AGA TCG AGT CCA TCC CGA GTA AAG GAG AAG AAC TTT TCA CTG

AO200 TAA GGA ATG GTA GAG TTT GAT AGG TTC CTA GCA GAT TAT TCG CTA CAG CAA ATT CTC AGT CCT GCT CCT CG

AO205 ATG AAG GCT CAT AGT GGT GC

JDpAL3A TTA TTG GTA CCC ATA CTA CTT TTC CGT CTG TCT TGC

JDpAL3C GCG TAT GGA TCC GGT AGA GTT TGA TAG GTT CCT AGC

JLNIPCA TTA TTG GTA CCG TTA TGG ATA ATC CTC TTG GAT ACG CTG ______

77 Strain Name Genotype ______

CB44 pyroA4; pyrG89 + alcA:tinC-HA pyr4+; wA3

CB45 pyroA4; pyrG89; + alcA:∆5 ’tinC-HA pyr4+; wA3

CDS165 GFP-sonB; pyroA4; pyrG89; wA3

CDS171 sonA-GFP; pyroA4; pyrG89; wA3

D23C Diploid between SO230 and SO223

GR5 pyroA4; pyrG89; wA3

JD70 pyroA4; pyrG89 + alcA::tinD-HA pyr4+; wA3

JD73 pyroA4; pyrG89 + alcA::∆5’ tinD-HA pyr4+; wA3

JD82 pyroA4; pyrG89 + ∆tinC pyr4+; wA3

JD83 pyroA4; pyrG89 + pyr4+; wA3

JD93 pyroA4 + pyroA+; pyrG89; wA3

JD98 pyroA4 + ∆An-hetC pyroA+; pyrG89; wA3

JD127 pyroA4 + ∆An-hetC pyroA+; pyrG89 + ∆tinC pyr4+; nicA2; chaA1

JD148 nimT23; pyrG89 + alcA: pyr4+; pabaA1; chaA1

JD154 nimT23; pyrG89 + alcA::∆5’tinC-HA pyr4+; pabaA1; chaA1

JD155 nimT23; pyrG89 + alcA:∆5’tinC-HA pyr4+; pabaA1; chaA1

JD157 GFP-sonB; pyroA4; pyrG89 + alcA: pyr4+; wA3

Continued

Table 2.2. List of A. nidulans strains.

78 Table 2.2. continued

JD158 GFP-sonB; pyroA4; pyrG89 + alcA::∆5’tinC-HA pyr4+; wA3

JD159 GFP-sonB; pyroA4; pyrG89 + alcA::∆5’tinD-HA pyr4+; wA3

JD160 sonA-GFP; pyroA4 + ∆An-hetC pyroA+; pyrG89 + ∆tinC pyr4+; chaA1

JD161 Diploid between strains SO221 and JD154

JD162 Diploid between strains SO221 and JD165

JD163 pyrG89 + tinC-GFP pyr4+; pyroA4; wA3

JD164 nimT23; pyrG89 + tinD-GFP pyr4+; pabaA1; chaA1

JD165 nimT23; pyrG89 + alcA:∆5’ tinD-HA pyr4+; pabaA1; chaA1

JD166 GFP-tubA; pyrG89 + alcA:∆5’ tinD-HA pyr4+; pabaA1; choA1; fw

LO1029 GFP-tubA; pyrG89; pabaA1; choA1; fw

R153 pyroA4; wA3

SO117 nimA7; pyrG89; pyroA4; nicA2; wA3

SO182 nimT23; pyrG89; pabaA1; chaA1

SO221 nimT23; pyrG89 + alcA:nimA(5C) pyr4+; nicA2; fwA1

SO223 nimT23; pyrG89 + alcA:nimA(5C) pyr4+; pabaA1; fwA1

SO230 nimT23; pyrG89 + alcA: pyr4+; pyroA1; chaA1

SWJ298 pyrG89; nicA2; chaA1 ______

79

CHAPTER 3

SEQUENCE CHARACTERIZATION AND SUBCELLULAR LOCALIZATION OF

TINC AND TIND

3.1. Introduction

The NIMA protein kinase plays an essential role in mitotic entry in A. nidulans. Also, a gathering body of evidence has identified the NIMA-like kinase

family members of other organisms as important cell cycle regulators. As a

protein kinase, NIMA is believed to interact with other proteins to regulate cell cycle progression. The identification of these NIMA substrates and interacting proteins will provide insight into the mechanisms through which NIMA acts and is acted upon in coordinating mitosis. At the time this study was initiated no NIMA targets or interacting proteins had been identified. Subsequent studies in the lab have identified histone H3 (De Souza et al., 2000) as a NIMA substrate, while two nucleoporins have been identified as NIMA interacting proteins (Wu et al.,

1998; De Souza et al., 2003). Additionally, the novel spindle pole body associated protein, TINA, was isolated in the initial Two-hybrid screen described here (Osmani et al., 2003). In addition to TINA, the Two-hybrid screen identified additional NIMA interacting proteins, including TINC and TIND. This introduction

80 describes the use of the yeast Two-hybrid screen to identify six potential NIMA

interacting proteins. General transformation characteristics of A. nidulans, the

exploitation of the auxotrophic marker pyrG89 for transformation, and an A. nidulans specific inducible expression system are also introduced.

3.1.1. The yeast Two-hybrid system.

The yeast Two-hybrid system provides a powerful system for the identification of protein-protein interactions. The system has been widely used since its inception due to ability to rapidly identify and recover genes which code for proteins that interact with a “bait” protein of interest (Fields and Song, 1989)

The Two-hybrid system is based on the ability to separate the GAL4 transcriptional activator into two functional domains: an Activation Domain (AD) and a DNA Binding Domain (BD) (Ma and Ptashne, 1987). The screen makes use of two different types of plasmids. The “bait” plasmid consists of bait gene sequence (in this case nimA) downstream of and in frame with the GAL4 BD sequence. The “target“ plasmids are produced by the “cloning” of a library of cDNAs downstream of and in frame with the GAL4 AD sequence.

Transformation of these plasmids in yeast cells results in the expression of two different types of fusion proteins: GAL4 BD-bait fusion proteins and GAL4 AD- target fusion proteins. The GAL4 BD fusion protein binds to specific GAL4 upstream activation sequences (UAS) in the yeast genome, however, the BD alone is insufficient to turn on transcription of downstream reporter genes, β- galactosidase (lacZ) and histidine (HIS3). A protein interaction between the bait

81 protein and the target protein brings the BD and AD into close enough proximity to reconstitute GAL4 transcriptional activation activity and allows for expression of downstream reporter genes. Protein interactions are identified in co- transformed yeast colonies expressing both target and bait fusion proteins by assaying for transcriptional activation of reporter genes.

3.1.2. NIMA interacting protein screen.

A Two-hybrid screen utilizing different versions of NIMA as bait was conducted in our laboratory by Aysha Osmani resulting in the identification of six

NIMA interacting proteins (TINA, B, C, D, E, F) (Osmani et al., 2003). GAL4 BD- nimA constructs were created and cotransformed into yeast with GAL4 AD constructs produced from a library of A. nidulans (Figure 3.1A).

Five different NIMA bait constructs were utilized in the screen (Figure

3.1B). The bait constructs included three kinase negative versions of NIMA: two which consisted of point mutations of essential residues within the NIMA kinase domain (K40M, and T199A) and a construct lacking a portion of the kinase domain (∆5’ & ∆3’). Two kinase active versions of NIMA (Full-length and ∆3’) were also used in the screen (Figure 3.1B).

The NIMA interacting proteins displayed varying levels of interaction and different specificities in terms of the NIMA baits with which they interacted. TINA interacted strongly with all versions of NIMA. In contrast, TINC and TIND displayed very selective affinity for NIMA, in that TINC interacted weakly with kinase inactive forms of NIMA, but failed to interact with kinase active forms of

82 NIMA. Finally, TIND interacted specifically with kinase active forms of NIMA but failed to interact with kinase inactive NIMA.

cDNAs corresponding to all six NIMA interacting proteins identified in the

Two-hybrid screen were expressed in A. nidulans. Overexpression of both TINC and TIND inhibited colony formation of A. nidulans. Examination of nuclear morphology in these cells revealed an underlying cell cycle defect. These cells exhibited polarized growth, albeit at a reduced rate compared to control cells, with an apparent lack of nuclear division. Nuclei in these cells were generally very large with uncondensed DNA and visible nucleoli, reminiscent of late interphase nuclei in control cells. In some cases these nuclei appeared highly stretched and smaller masses of DNA were observed separate from the main mass of DNA. The identification of TINC and TIND as NIMA interacting proteins coupled with the ability of these proteins to induce nuclear division defects prompted further investigation of the roles of these proteins in A. nidulans

To begin to investigate the functions of TINC and TIND and their potential roles in cell cycle regulation we sought to clone tinC and tinD to determine whether they had been characterized in A. nidulans or whether potential homologues of known function existed in other organisms. Additionally regulated expression of TINC and TIND was used to examine their localization in A. nidulans. Considering the function of NIMA in A. nidulans we expected that the

NIMA interacting proteins identified in the Two-hybrid screen may represent spindle components, DNA accessory proteins including histones or condensins, or nuclear pore components. The massive sequencing projects conducted

83 across a broad range of organisms has provided a wealth of sequence information and the ability to quickly identify potential protein homologies in an organism or across species. This chapter describes the sequence characterizations for TINC and TIND, as well, as the subcellular localization of these proteins in A. nidulans.

TINC and TIND sequence analysis revealed surprising identities. TINC shared a high degree of homology with Het-C, a N. crassa protein involved in heterokaryon incompatibility. TIND is a highly conserved protein with homologues involved in Fe-S cluster biogenesis. I will present a brief introduction to these topics as well as a description of the alcA promoter based system used for regulated gene exspression in A. nidulans.

3.1.3. Expression proteins in A. nidulans using the alcA promoter.

Proteins of interest can be expressed in A. nidulans using a plasmid system based on the promoter from the alcohol dehydrogenase 1 gene (Waring et al., 1989). The plasmids used for these expression studies were pAL5 and pAL3 (Figure 3.2). pAL3 is based on pUC19 and includes N. crassa pyr-4, as well as, a multi-cloning site immediately downstream of the promoter for the alcA

(encodes alcohol dehydrogenase I) gene (Waring et al., 1989). Genes inserted at this cloning site were demonstrated to be under the control of the alcA promoter at the level of transcription when introduced into A. nidulans (Waring et al., 1989). alcA based promoter expression is known to be greatly repressed, at the level of transcription, in the presence of glucose (Bailey and Arst, Jr., 1975;

84 Lockington et al., 1985; Lockington et al., 1987) and markedly induced in the presence of ethanol (Creaser et al., 1985). Glycerol was used as a non-inducing, non-repressing carbon source. alcA expression can be further varied in liquid cultures grown in minimal yeast lactose media by addition of different concentrations of threonine.

The pAL5 plasmid was generated by adding a 3.8 Kb fragment of H2A flanking sequence, positioned downstream of the pAL5 cloning site, to assist in site-specific recombination and 3’ processing of cDNA inserts (Figure 3.2).

3.1.4. Heterokaryon incompatibility

A heterokaryon is defined as an organism containing two or more genetically unlike nuclei. It has long been recognized that filamentous fungi are capable of producing heterokaryons through a process of hyphal fusion between two “donor” organisms. The result of this hyphal fusion produces a fungal heterokaryon containing two populations of nuclei which are mixed along the length of the hyphal segments of the organism. While the ability of some fungi to produce heterokaryons, A. nidulans included, represents a valuable genetic research tool, the reasons for their occurrence in wild isolates of filamentous fungi is not well understood.

Not all strains within a species of filamentous fungi are capable of forming a viable heterokaryon with every other strain within that same species. This restriction on heterokaryon formation between genetically dissimilar organisms has been termed vegetative or heterokaryon incompatibility and is thought to be

85 important in the maintenance of organism identity and the prevention of virus

transmission (Caten, 1972; Todd and Rayner, 1980). Heterokaryon

incompatibility was first identified in A. nidulans in 1963 (Grindle, 1963a; Jinks

and Grindle, 1963; Grindle, 1963b), and is known to exist in other fungi, including

N. crassa.

Heterokaryon compatibility between filamentous fungi is determined by a

class of allelic nuclear genes termed either het (heterokaryon incompatibility

gene) or vic (vegetative incompatibility gene) such that only organisms with

compatible het loci are capable of forming heterokaryons with each other. At

least eight het loci have been identified in A. nidulans, some of which are known

to be multi-allelic (Anwar et al., 1993; Dales et al., 1993). In N. crassa, at least

ten het genes are known to function (Perkins, 1988). Strains with incompatible

het genes are able to undergo hyphal fusion but the heterokaryotic hyphae are

rapidly destroyed by the initiation of an incompatibility response similar to

programmed cell death (Garnjobst and Wilson, 1956; Jinks and Grindle, 1963;

Mylyk, 1975; Jacobson et al., 1998; Marek et al., 2003).

het-c is the most thoroughly characterized het locus in N. crassa. The

locus is known to be multi-allelic with alleles designated het-COR, het-CEM, and het-CPA (Garnjobst and Wilson, 1956; Howlett et al., 1993). Het-C contains an N-

terminal consensus signal peptide and a coiled-coiled domain between residues

426 to 458. Analysis of forms of Het-C which lack this coiled coil region result in

a protein less able to initiate incompatibility reactions, suggesting that this coiled-

coil domain may play a role in mediating the incompatibility response through

86 protein-protein interactions (Saupe et al., 1996). Deletion studies also identified

the region between residues 581 to 729 as essential in triggering the het-C

incompatibility response (Saupe et al., 1996). Sequence manipulation of this

region produces forms of Het-C with differing incompatibility specificity (Wu and

Glass, 2001).

3.1.5. Fe-S clusters

Fe-S clusters are comprised of inorganic iron and sulfur coordinated by

cysteines within a protein. Fe-S clusters are common prosthetic groups of

proteins involved in electron transport. Fe-S assembly is facilitated by a family of

proteins first described in the nitrogen fixing bacteria Azotobacter vinelandii

(Yuvaniyama et al., 2000). NIFU was identified as one of the proteins involved in

coordinating Fe-S clusters for assembly into nitrogenase, a key enzyme involved

in the reduction of atmospheric nitrogen (Burgess and Lowe, 1996; Yuvaniyama

et al., 2000). NIFU is a modular protein which is thought to act as a scaffold

protein for Fe-S assembly (Agar et al., 2000). Each module is characterized by

the presence of cysteines involved in coordinating assembly. The amino-

terminal module is termed the Transient Cluster Module since Fe-S clusters

assembled here will be transferred to target proteins (Agar et al., 2000;

Yuvaniyama et al., 2000). The middle module, or Permanent Cluster Module,

contains an Fe-S cluster which remains associated with NIFU and which is

required for its function. Finally, the function of the carboxy-terminal domain of

NIFU is undetermined, however, it has been termed the Thioredoxin Module due

87 to the fact that the two cysteines present within this module exist in a similar sequence context to those found in the catalytic site of thioredoxin (Holmgren,

1985; Frazzon et al., 2002). In organisms other than bacteria these modules are split between two or more proteins. For example, in the yeast S. cerevisiae the proteins ISU1 an ISU2 contain a domain which is homologous to the amino- terminal domain of NIFU, while the protein NFU1 contains a domain which is homologous to the carboxy-terminus of NIFU (Schilke et al., 1999). NFU1 is highly conserved as homologues have been identified in humans, mice and A. nidulans (TIND), all of which contain the carboxy-terminal module of NIFU

(Lorain et al., 2001; Ganesh et al., 2003).

3.2. Results

3.2.1. tinC cloning and sequence characterization.

The plasmid containing the tinC cDNA was recovered from the Two-hybrid screen and sequenced. Sequencing analysis identified an open reading frame of just over 2.5 kb (2,536 base pairs). Since construction of library cDNA “target” plasmids for the Two-hybrid screen results in an in-frame fusion between the 5’ end of the cDNA and the 3’ end of the GAL4 Activation Domain, it was thought that cDNA which was recovered may not contain the entire tinC coding region. A full-length tinC cDNA was identified by 5’ RACE PCR. Sequencing analysis of the tinC cDNA clone identified a 2,937 open reading frame, 397 base pairs longer than the open reading frame sequenced for the original 5’ truncated

88 tinC cDNA. The open reading frame encodes a protein of 978 amino acids with a theoretical molecular weight of 110 kD (Figure 3.3A).

Sequence comparisons between the full-length tinC cDNA clone obtained from 5’ RACE PCR and the original tinC cDNA identified from the Two-hybrid screen identified three point mutations in the full-length cDNA sequence. These mutations are believed to be due to the rounds of PCR which occurred prior to sequencing. Due to the nature of the reaction a Taq based non-proofreading polymerase mix must be used for 5’ RACE PCR applications.

PCR screening of a sorted A. nidulans cosmid library, identified chromosome six as the location of the tinC gene. Sequence comparisons between tinC sequence obtained from the tinC containing cosmid and the tinC cDNA identified three introns of forty-five base pairs, forty-seven base pairs, and sixty-two base pairs within the coding region (Figure 3.3B). All three introns conformed to the consensus intron start and stop sequences with each intron beginning with GT and ending with AG. Alignment of gtinC to the tinC cDNA sequence also confirmed the presence of three point mutations within the tinC cDNA. Therefore, this tinC cDNA clone was not used in any downstream analysis.

Sequence analysis of the projected TINC protein revealed several potential functional motifs. First, the consensus NIMA phosphorylation site has been defined as a serine or threonine which is three residues downstream from a phenylalanine (FxxT/S) (Lu et al., 1994). TINC contains three sites which match this NIMA consensus phosphorylation site (Figure 3.3A). Note that the

89 phosphorylation site nearest the amino-terminus of TINC would not be present in

amino truncated versions of TINC from the Two-hybrid screen. . Second, TINC also contains a potential coiled-coil domain between amino acids 250 to 400

(Figure 3.3A). Finally, the amino-terminus of TINC contains a predicted signal peptide (Figure 3.3A). While signal peptides display relatively little primary sequence conservation they consistently contain three domains termed “c”, “h” and “n” of relatively consistent size and residue composition (Ma and Ptashne,

1987). The c-region terminates at the peptide cleavage site (Ma and Ptashne,

1987). The h-region is hydrophobic and consists of seven to thirteen residues

(Bird and Bradshaw, 1997). The length of the n-region is variable, but always contains a net positive charge. Analysis of TINC sequence using the pSORT program demonstrated that the amino-terminus of TINC correlated well with these requirements and identified a potential cleavage site between amino acid residues twenty-four and twenty-five (Ballance and Turner, 1985).

3.2.2. TINC is a member of a fungal specific family of proteins

The predicted TINC protein sequence was used to perform BLASTp homology searches at NCBI. Initial searches identified a protein from N. crassa,

Het-C, which shared a high level of homology with TINC. Het-C is known to be involved in regulating heterokaryon incompatibility and programmed cell death in

N. crassa. Global comparative alignments of the two sequences using ClustalW showed them to be thirty-nine percent identical and sixty-three percent similar.

90 Additional BLASTp homology searches performed at both NCBI and the

Whitehead Institute identified additional TINC-related proteins in other filamentous fungi and in A. nidulans itself (Figure 3.4). BLAST homology searches in an A. nidulans sequence database (The Whitehead Institute) identified an uncharacterized potential protein, AN2167.2, which is 39.5% identical to TINC (Figure 3.4). We have named AN2167.2 An-HETC due to high level of sequence identity (48.3%) it shares with the N. crassa protein Het-C

(Figure 3.4). Note that the degree of sequence identity between An-HET-C and

Het-C (48.3%) is greater than the degree of identity between TINC and Het-C

(39%). BLASTp homology searches performed using the sequence databases of other filamentous fungi identified additional proteins which share significant homology with TINC. Specifically, three TINC-related predicted proteins were identified in Fusarium graminearum (FG00728.1, FG05163.1, and FG02905.1) and Magnaporthe grisea (MG01326.4, MG09383.4, and MG03919.4) (Figure 3.4).

Analysis of ClustalW sequence alignments of the identified TINC-related proteins indicated that filamentous fungi contain a family of related proteins consisting of two main evolutionary branches, a TINC-like branch and a Het-C-like branch

(Figure 3.4).

To investigate whether TINC or An-TINC may play a role in heterokaryon incompatibility in A. nidulans, twenty-five A. nidulans strain isolates, representing twenty-one heterokaryon compatibility groups (Dr. Fons Debets, personal communication, Dales and Croft, 1990; Anwar et al., 1993), were sequenced at the tinC and An-hetC loci. In N. crassa Het-C is a multi-allelic loci (Saupe et al.,

91 1996). A polymorphic sequence region within Het-C has been shown to be

responsible for determining heterokaryon incompatibility (Saupe and Glass,

1997). Sequencing analysis identified no evidence for multi-allelism either at the

tinC or the An-hetC loci. Sequence analysis of An-hetC did identify four sequence dissimilarities, consisting of single nucleotide changes, within strains when compared to a parental Glasgow isolate strain. Three of these four mutations were silent, while the fourth mutation resulted in the use of leucine in place of phenylalanine. Both of these amino acids are neutral and non-polar in character.

3.2.3. TINC antibody characterization

A peptide antibody (α-TINC) was generated to the TINC specific amino

acid sequence KGEYEPQGYERQGSQL. Total protein lysates were generated

from an asynchronous A. nidulans culture, and subjected to SDS-PAGE. Total

protein lysateswere subjected to western blot analysis using the α-TINC antibody.

The antibody specifically recognizes a single protein which migrates slightly

slower than a 97 kD protein marker (Figure 3.5, lane 1). Furthermore, α-TINC

binding was efficiently competed by addition of a TINC-specific peptide (Figure

3.5, lane 2), but a non-specific peptide fails to compete (Figure 3.5, lane 3).

Additionally, the α-TINC antibody was able to recognize HA-epitope tagged forms

of TINC which were expressed in A. nidulans, further confirming that the protein

recognized by α-TINC is indeed TINC.

92 3.2.4. TINC localization

As described previously, predictions of TINC localization using the

PSORTII subcellular localization prediction program identified an N-terminal signal peptide, which should direct TINC into the secretory pathway. To investigate TINC localization in A. nidulans, a construct was produced which allows for expression of a version of TINC containing three hemagglutinin (HA) repeats at its carboxy terminus under the control of the alcA promoter. This construct was transformed into A. nidulans. The presence of TINC-HA in transformants was confirmed by western blotting of total protein lysates prepared from transformant cultures induced to express TINC-HA by germination in the presence of 40mM threonine. To examine TINC-HA localization, A. nidulans strains transformed with the alcA::tinC-HA construct were germinated on coverslips in minimal media glycerol. Glycerol was used as a carbon source rather than ethanol for these experiments to attempt to decrease the chance for localization artifacts due to TINC overexpression. Glycerol induces a much lower level of alcA promoter driven expression than ethanol. After allowing cells to germinate, cells were fixed and processed for immunofluorescence using anti-HA antibodies. Examination of TINC-HA by immunofluorescence microscopy revealed the presence of TINC-HA throughout the cytoplasm while some areas lacked TINC-HA staining (Data not shown). DAPI staining revealed that areas lacking TINC-HA staining corresponded to nuclei (Data not shown). A TINC-GFP fusion construct was created which allowed for the expression of TINC-GFP under the regulation of the tinC promoter to examine TINC localization in living

93 cells. A. nidulans transformants containing the tinC-GFP construct were screened for TINC-GFP expression by microscopic confirmation of GFP fluorescence and western blotting of transformant total cell lysates. Living cells expressing TINC-GFP (JD163) were examined using confocal microscopy.

TINC-GFP was observed throughout the cell cytoplasm (Figure 3.6A). Fixation and DAPI of these cells revealed that areas lacking TINC-GFP corresponded to nuclei (Figure 3.6B). To confirm the cytoplasmic localization of endogenous

TINC, TINC localization was examined using the α-TINC antibody. A. nidulans strain R153 was germinated in YG media at 32°C. This strain was fixed and processed for immunofluorescence using α-TINC antibodies. As described for

TINC-HA and TINC-GFP, TINC was present throughout the cytoplasm in a punctuate pattern (Figure 3.6C). TINC was excluded from the nuclei of most cells. Note that the cell depicted in Figure 3.6C is a projection of vertical image slices through a single cell, as such, the nuclei are more difficult to discern than in a single image slice.

Finally, as mentioned, a small portion of all cells examined for TINC

localization (~5%) displayed continuous TINC throughout the entire cell. This

pattern was suggestive of the fact that these cells may be mitotic. To examine

this possibility, cells expressing TINC-GFP were fixed and processed for

immunofluorescence utilizing an anti-tubulin antibody. The presence of mitotic

spindles and condensed DNA indicated that cells were in mitosis. Examination

of these cells by indirect immunofluorescence microscopy (Figure 3.6D)

confirmed that (a) interphase cells contained areas of negative fluorescence

94 corresponding to nuclei, while (b) mitotic cells contained TINC GFP both in the

cytoplasm and nuclei. Therefore TINC is a cytoplasmic protein throughout interphase and is present in nuclei during mitosis.

3.2.5. tinD cloning and sequence characterization

The plasmid containing the tinD cDNA was recovered from the Two-hybrid screen and sequenced. Sequencing analysis identified an open reading frame of

768 bp. The first potential initiation methionine did not occur until 449 nucleotides into the sequence suggesting that this cDNA was almost certainly 5’ truncated. A full length tinD cDNA was obtained by performing 5’ RACE PCR.

Sequencing analysis of the full-length tinD cDNA clone identified a 981 base pair open reading frame. The open reading frame encodes a protein of 326 amino acids with a theoretical molecular weight of 35.8 kD (Figure 3.7A).

A genomic tinD clone was isolated in the same manner described for tinC. tinD was determined to lie on chromosome 8. Sequence comparisons between tinD and the tinD cDNA identified two introns of sixty-four base pairs and seventy base pairs within the coding region (Figure 3.7B). Both introns conformed to the consensus intron start and stop sequences with each intron beginning with GT and ending with AG.

Sequence analysis of the projected TIND protein revealed several potential functional motifs. First, TIND contains three sites which match the

NIMA consensus phosphorylation site (Figure 3.7A). Second, TIND also contains a potential coiled coil domain between amino acids 275 to 295 (Figure

95 3.7A). Third, the amino-terminus of TIND contains a mitochondrial targeting

peptide with a projected cleavage site at residue 79 (predicted by MiTOP) or 97

(Predicted by pSORTII) depending on the prediction program (Figure 3.7A).

Finally, the carboxy-terminus of TIND contains a region with a high degree of

homology to the carboxy-terminal domain of the bacterial NIFU protein (Figure

3.7A).

3.2.6. TIND is a highly conserved protein

The predicted TIND protein sequence was used to perform BLASTp

homology searches at NCBI. Initial searches identified proteins in a range of

organisms from bacteria and yeast through to humans. Sequence alignments using ClustalW between TIND homologues indicated that TIND was 35.5% identical to S. cerevisiae Nfu1, 42.1% identical to a human TIND homologue and

48.4% identical to a Rickettsia prowazekii TIND homologue (Figure 3.8A).

Homologues displayed the highest degree of identity in their carboxy terminus in which they all contained a domain homologous to the NIFU carboxy-terminal domain NIFU protein module (Figure 3.8B). NIFU is a modular protein involved in Fe-S cluster assembly. NIFU was first characterized in nitrogen fixing bacteria.

Figure 3.8C shows an alignment between the carboxy terminus of TIND (An) and the carboxy terminus of NIFU from the nitrogen fixing bacteria Anabaena azollae

(Aa). The ** indicate a pair of conserved cysteines which are conserved in a context similar to that of thioredoxin (Figure 3.8C).

96 3.2.7. TIND localization

Predictions of TIND localization using the PSORTII subcellular localization

prediction program identified an N-terminal mitochondrial targeting peptide. To

investigate TIND localization in A. nidulans, TIND-HA was expressed in A.

nidulans under the control of the alcA promoter. TIND localization was examined in a strain expressing TIND-HA (JD70) by immunofluorescence and anti-HA antibodies. Examination of TIND-HA by immunofluorescence microscopy

revealed the presence of TIND-HA in filamentous organelles (Figure 3.9A) which

colocalized with the mitochondrial marker MitoTracker (Molecular Probes)

(Figure 3.9B). In order to examine TIND localization in living cells, a tinD-GFP

fusion construct was created which allowed for the expression of TIND-GFP

under the regulation of the tinD promoter. Living cells which expressed TIND-

GFP (JD164) were examined using confocal microscopy. In fixed cells, TIND-

GFP was observed in filamentous structures which colocalized with the mitochondrial marker MitoTracker (Figure 3.9C and D). In living cells TIND-GFP also appeared to localize exclusively to highly filamentous organelles, which were shown to be mitochondria by MitoTracker staining (Figure 3.9E).

To determine whether TIND localization changes through the cell cycle,

cells expressing TIND-GFP were costained with anti-tubulin antibodies and DAPI

staining. Examination of these cells by immunofluorescence microscopy showed

that TIND was present in the mitochondria throughout the cell cycle, during

interphase (Figure 3.10A) and mitosis (Figure 3.10B and C).

97 3.2.8. Identification of human tinD cDNAs

At the time these studies were conducted, a full-length human TIND cDNA had not been isolated, therefore we undertook to identify a full-length cDNA and to determine whether human cells contain potentially non-mitochondrial forms of

TIND. To this end 5’RACE PCR was conducted using primer AO153 in a

Marathon Ready HeLa cDNA library (Clontech). Intriguingly, the cDNAs isolated in this screen could be grouped into three main transcripts (Figure 3.11). Two classes of cDNAs were predicted to encode mitochondrial forms of human TIND

(Figure 3.11A and B). Notably, the majority of cDNAs isolated lacked the second in-frame methionine due to a T to A change at this position (Figure 3.11A).

These cDNAS were significantly longer than those reported at the time. A third class of cDNAs contained all three in-frame methionines, however, a 63 bp insert between the first and second methionine contained an in-frame stop. This opened the possibility that translation of this transcript could produce non- mitochondrial forms of human TIND by initiating at the second methionine.

3.3. Discussion

3.3.1. TINC and heterokaryon incompatibility

The high degree of homology between TINC and Het-C initially indicated a role for TINC in regulating heterokaryon incompatibility; however, several additional pieces of information make this unlikely. First, the availability of the complete genome sequence allowed for the identification of An-HETC, based on

98 TINC homology searches. Sequence comparisons between TINC, An-HET-C

and Het-C revealed that An-HETC is more similar to Het-C than is TINC. Second,

sequence analysis of tinC and An-hetC from twenty-five A. nidulans strains

representing twenty-one different incompatibility types revealed no evidence for

multi-allelism at either of these loci, a defining characteristic of compatibility

determination at the het-C locus of N. crassa (Saupe and Glass, 1997; Wu and

Glass, 2001). Third, characterization of a Podospora anserina het-C homologue,

hch, failed to identify evidence for multi-allelism at this loci and suggested that

hch may not be involved in heterokaryon incompatibility (Saupe et al., 2000).

These results suggest that some proteins in N. crassa may have evolved

incompatibility functions which are species specific.

The presence of the tinC-related gene An-hetC, and the conservation of

two such genes in additional filamentous fungi, suggests that TINC plays an

important function in A. nidulans. Our early overexpression data and NIMA

interaction data suggest that TINC may be involved in cell cycle regulation in A.

nidulans. This possibility will be analyzed by generating deletion mutants of tinC,

further characterization of the tinC cell cycle specific overexpression defect, and attempts to co-immunoprecipitate NIMA and TINC in A. nidulans.

3.3.2. TIND and Fe-S cluster assembly

BLAST homology searches conducted using TIND identified TIND

sequence homologues in organisms ranging from bacteria through to humans.

All homologues contained a highly conserved region which was highly similar to

99 the carboxy-terminal domain of NIFU, a bacterial protein originally identified for its role in nitrogen fixation. NIFU has been implicated as acting as a scaffold protein to generate Fe-S clusters for insertion into nitrogenase. Significantly, only the carboxy-terminal domain is present in TIND, and while the two amino terminal domains of NIFU have defined roles in Fe-S cluster assembly the function of the carboxy-terminal domain remain unknown. Studies in yeast have suggested a potential role for Nfu1 in Fe-S biogenesis and iron metabolism, as

∆nfu1 ∆isu1 cells display decreased mitochondrial respiration activity and accumulate iron (Schilke et al., 1999). This domain does possess two highly conserved cysteines which occur in a context similar to the of thioredoxin, an enzyme involved in redox regulation. The potential significance of this relationship will be discussed in Chapter 7.

3.3.3. TINC localization

The presence of a potential amino-terminal signal peptide in TINC suggested that TINC would be directed to enter the secretory network in A. nidulans. Previous studies in Aspergillus have identified the hyphal tips as primary sites of protein secretion (Wosten et al., 1991). Secreted proteins frequently show accumulation at hyphal tips and septa along the length of the hyphae (Gordon et al., 2000; Khalaj et al., 2001). Given this information, the presence of TINC in the cytoplasm was somewhat surprising. The use of two forms of epitope tagged TINC and examination of endogenous TINC localization using a TINC specific antibody argue that TINC is a cytoplasmic protein. Such a

100 localization pattern suggests that the signal peptide in TINC may be non-

functional. Interestingly, studies of heterokaryon incompatibility in N. crassa involving plasmid transformation have demonstrated that HET-C function is independent of the signal peptide (Louise Glass personal communication). Het-

C localization studies also failed to show Het-C at hyphal tips or cell wall (Sarkar et al., 2002). Although, our studies suggest that TINC does not participate in heterokaryon incompatibility reactions, the high degree of sequence similarity it shares with Het-C make these Het-C studies relevant when considering TINC localization. Finally, the localization of TINC to nuclei during mitosis also suggests that NIMA and TINC may interact during mitosis.

3.3.4. TIND localization

Studies in S. cerevisiae had shown that the TIND homologue, Nfu1, was mitochondrial (Schilke et al., 1999). First, the authors of the study identified

NFU1 based on its genetic interaction with SSQ1, a mitochondrial heat shock protein. Second, Nfu1 contained a mitochondrial targeting peptide. Third, they demonstrated that radiolabeled Nfu1 was efficiently taken up and processed by active mitochondria. Fourth, mitochondrial purification and fractionation performed in a strain which expressed myc-tagged forms of Nfu1 demonstrated that Nfu1 was present in the mitochondrial matrix (Schilke et al., 1999).

Therefore, the localization of both TIND-HA and TIND-GFP to mitochondria in A. nidulans was not surprising considering the Nfu1 localization data and given the fact that TIND possessed a predicted amino-terminal mitochondrial targeting

101 peptide. The fact that TIND was isolated as a NIMA interacting protein, however, is surprising given the known function of NIMA and previous NIMA localization studies.

During late G2, NIMA is present throughout the cytoplasm. At the G2-M transition NIMA is enriched at the nuclear periphery. During mitosis, NIMA is present in the nucleus, and frequently co-localizes with the mitotic spindle (De

Souza et al., 2000). We had hoped that TIND might be present outside the mitochondria during some phase of the cell cycle, particularly mitosis, to suggest a possible interaction with NIMA, but examination of TIND localization during mitosis and interphase argues against this idea. Importantly, these localization studies all rely on the expression of epitope tagged forms of TIND. The inability to produce a functional TIND specific antibody leaves open the possibility that epitope tagged versions of TIND may not reflect the localization of endogenous

TIND. Also, these studies do not fully rule out the possibility that a minor population of TIND, which is not discernable by fluorescence microscopy, is present outside of the mitochondria.

Additionally, conflicting reports on the localization of the human homologue of TIND, HIRIP5 suggest that TIND may be present outside of mitochondria. HIRIP5 was first identified in a Two-hybrid screen as a HIRA interacting protein (Lorain et al., 2001). HIRA acts as a transcriptional co- repressor with predominantly nuclear localization and is thought to be involved in the development of DiGeorge syndrome (Lamour et al., 1995; Lorain et al., 1998;

Magnaghi et al., 1998). The cDNA isolated by the authors of this study was 5’

102 truncated leading them to propose multiple translation initiation sites and suggest

a potential nuclear role for HIRIP5 (Lorain et al., 2001). In a second study,

HIRIP5 was identified as a laforin interacting protein with a cytoplasmic localization (Ganesh et al., 2003). This study identified a full-length HIRIP5 cDNA, analogous to those which I isolated. This cDNA extended the predicted

HIRIP5 protein by twenty five amino acids. Localization and fractionation experiments performed in this study showed that HIRIP5 was a cytoplasmic protein (Ganesh et al., 2003) suggesting that the mitochondrial targeting peptide

in HIRIP5 is not functional or that this cytoplasmic form of HIRIP5 may represent

an alternately spliced form. Finally, the most recent study of the localization of

the human homologue of TIND identified two different transcripts analogous to

those which I identified (Tong et al., 2003). Isoforms which contained the first

methione were present in the mitochondria, while a shorter form which initiates

from the second in-frame methionine is found in the cytoplasm and nucleus

(Tong et al., 2003).

There is evidence for the presence of additional Fe-S assembly machinery

in non-mitochondrial compartments. The human Nifs protein, a cysteine

desulfurase proposed to supply the inorganic sulfur in iron-sulfur clusters, is

found in the mitochondria, cytoplasm and nucleus (Land and Rouault, 1998).

These different forms are produced from a single nifS transcript through different in-fame start methionines. Initiation was regulated by the pH of the cytosol and media (Land and Rouault, 1998). This study prompted the search for alternate

TIND cDNAs in A. nidulans which may encode forms of TIND localized to

103 different cellular compartments, however, all cDNAs isolated encoded

mitochondria directed TIND. The presence of several in-frame methionines in

TIND leaves open the possibility that under some situations, including differing

pH conditions, different forms of TIND are produced.

Finally, it remains a formal possibility that TIND exists outside the

mitochondria under cell growth conditions which were not examined. An

intriguing possibility is that TIND may be released from the mitochondria during

apoptosis. Studies have demonstrated that the addition of sphingoid long-chain

bases to A. nidulans induces programmed cell death (Cheng et al., 2003).

Preliminary studies I performed using these sphingolipids suggest that TIND

remains within mitochondria even after several hours of treatment (Data not

shown). It will be important to determine whether TIND and NIMA interact with

each other in A nidulans through co-immunoprecipitation studies. Alternately, it

may prove that NIMA and TIND do not interact in A. nidulans. If so, it may be that TIND was isolated as a NIMA Two-hybrid interacting protein due to its artificial localization in this screen. This second possibility is worth further study, however, due to the ability of ∆N-TIND to interact with NIMA in yeast and the ability of ∆N-TIND expression to produce a cell cycle specific defect strikingly similar to a nim defect. It will be important to investigate whether full-length TIND expression can induce cell cycle defects in A. nidulans.

104 A Z X Y NIMA AD AD AD BD lacZ gene (HIS-) NIMA TIN BD AD lacZ gene (HIS+) B +ve Full-length Kinase Catalytic Domain 3'

5' & 3' K40M -ve K40M Kinase T199A T199A Kinase-ve

Figure 3.1. Identification of NIMA interating proteins using the yeast Two-

hybrid system. A yeast two hybrid screen was used to identify NIMA interacting proteins. (A) Fusion proteins were created beween NIMA and the DNA Binding

Domain of GAL4 (BD). Fusion proteins were also generated between a library of

A. nidulans cDNAs and the Activation Domain of GAL4 (AD). An interaction between NIMA and a NIMA interacting protein (TIN) reconstituted the GAL4 transcriptional activator and turned on downstream reporter genes. (B) Five different NIMA constructs were used on the screen. Three versions of NIMA were kinase inactive (∆5’ & ∆ 3’, K40M, and T199A), while two versions of NIMA were active kinase constructs (Full-length and ∆3’).

105 M a CS pyr-4 lcA ( 2.3 Kb)

pAL5 8.86 Kb H2A 3' end (3.8 Kb)

pUC19 (2.7 Kb)

Figure 3.2. pAL5. Genes of interest can be cloned into pAL5 downstream from the alcA promoter. pAL5 also contains sequences from the 3’ end of H2A and the N. crassa pyr-4 gene which complements the auxotrophic pyrG89 mutation in

A. nidulans.

106 A Potential Potential NIMA signal phosphorylation sites (FXXT/S) peptide

P P P sP CC 110kD

Fusion point in Two-Hybrid Potential coiled-coil domain library

0 100 200 300 400 500 600 700 800 900 1000 (aa)

B ATG TAA 1 2 Exon 3 Exon 4

0 1000 2000 3000

Figure 3.3. TINC. (A) tinC encodes a protein of 978 amino acids with a theoretical molecular weight of 110 kD. TINC contains a predicted coile coil domain (CC), an amino-terminal signal peptide (sP), and three consensus NIMA phosphorylation sites (P). (B) tinC consists of four exons with intervening introns of 45 bp, 47 bp, and 62 bp.

107

Figure 3.4. TINC is a member of a fungal-specific family of proteins.

BLASTp homology searches using TINC identified a family of related proteins in

filamentous fungi, which includes the N. crassa Het-C protein and An-HETC in A. nidulans. A phylogenetic tree was generated following alignments of proteins by

ClustalW.

108

% N. crassa % TINC Het-C Identity Identity

(NCU03125.1) 49.6 34.5

(FG00728.1) 45.6 35.3

(MG01326.4) 52.3 34.8

(MG09383.4) 41.9 32.6 109

(AN9067.2) TINC N/A 34.6

(FG05163.1) 39.1 60.3

(NCU03493.1) Het-C 34.6 N/A

(MG03919.4) 37.6 57.1

(AN2167.2) An-HETC 39.5 48.3

(FG02905.1) 31.8 29.2

Lanes 132 α-TINC Ab + + + TINC Pep --+ Non-specific Pep - - +

97 kD

Figure 3.5. Characterization of a TINC specific antibody. Total protein lysates were prepared from A. nidulans strain R153 and subjected to SDS-PAGE and western blotting using α–TINC antibodies.

110

Figure 3.6. TINC localization. (A) A. nidulans transformants which expressed

TINC-GFP were germinated in minimal media. TINC-GFP localization was

examined in living cells using a spinning disk confocal microscopy system

(Perkin Elmer). (B) To visualize nuclei, cells were fixed, stained with DAPI and

examined using immunofluorescence microscopy. (C) A. nidulans strain GR5

was germinated in YG media at 32°C, fixed and processed for indirect

immunofluorescence microscopy. TINC localization was examined using α-TINC

antibodies followed by Alexa Fluor 488 conjugated goat anti-rabbit IgG

(Molecular Probes). (D) Cells expressing TINC-GFP were germinated in minimal media, fixed and processed for indirect immunofluorescence microscopy. DNA was stained with DAPI. Microtubules were visualized with mouse monoclonal antibody TAT-1 followed by AlexaFluor 594 conjugated rat anti-mouse IgG

(Molecular Probes). Cells were determined to be in (a) interphase or (b) mitosis based on DNA condensation and microtubule morphology.

111 A TINC-GFP

B TINC-GFP

DAPI

C α-TINC Ab

DAPI

D TINC-GFP a

b DAPI

Tubulin

112 A Coiled coil P P P mTP C-terminus NIFU

0 100 200 300 350aa

B ATG TAA Exon 1 Exon 2 Exon 3

0 1000 (Bp)

Figure 3.7 TIND. (A) tinD encodes a protein of 326 amino acids with a theoretical molecular weight of 35.8 kD. TIND contains a predicted coiled coil domain (CC), an amino-terminal mitochondrial targeting peptide (mTP), three consensus NIMA phosphorylation sites (P), and a carboxy-terminal domain homologous to the carboxy-terminal domain of the bacterial NIFU protein. (B) tinD consists of three exons

113 A A.nidulans MASSSTSANMVKRTILSNRLAAQILNFAEASPRTIRPFAFGSIRSIHNSARLPTITASET 60 H.sapiens MAATARRG------WGAAAVAAGLRRRFCHMLKNPYTIKKQP 36 S.cerevisiae ------MFKS 4 R.prowazekii ------

A.nidulans RHRPTLRP-HQLSAAHGRVNGQPPTGKRTIFIQTESTPNPDALKFIPNHRVLPEDFPTSF 119 H.sapiens LHQFVQRPLFPLPAAFYHP------VRYMFIQTQDTPNPNSLKFIPGKPVL----ETRT 85 S.cerevisiae VAKLGKSPIFYLNS------QRLIHIKTLTTPNENALKFLSTDGEMLQTRGSKS 52 R.prowazekii ------MFIQTEETPNPDAIKFFPGQEIS----VDQP 27 :.*:* *** :::**:. .

A.nidulans LEYLSPRSTLAPPHPSTLAANLFN-VEGVQSVFFGTDFITVTKASD-TNWAHIKPEVFSL 177 H.sapiens MDFPTPAAAF----RSPLARQLFR-IEGVKSVFFGPDFITVTKENEELDWNLLKPDIYAT 140 S.cerevisiae IVIKNTDENLIN--HSKLAQQIFLQCPGVESLMIGDDFLTIN-KDRMVHWNSIKPEIIDL 109 R.prowazekii VFFSELAEVKG---RSALAESLFH-INNVKSVFLGSDFITVTKQAR-GNWQVIKPEILMV 82 : * ** .:* .*:*:::* **:*:. .* :**::

A.nidulans ITQAVTSGEPIVNTVEKSGASGQKGG----EEDSLSYNEEDDEVVSMIKELLETRIRPAI 233 H.sapiens IMDFFASGLPLVTEETPSGEAGS------EED------DEVVAMIKELLDTRIRPTV 185 S.cerevisiae LTKQLAYGEDVISKEFHAVQEEEGEGGYKINMPKFELTEEDEEVSELIEELIDTRIRPAI 169 R.prowazekii IMDHFISGFPVFNENTKIDDEKH------NLDML------SEIEKQIIETIETRVRPFV 129 : . . * :.. : .*: * * ::**:** :

A.nidulans QEDGGDIEFRGFEN--GIVMLKLRGACRTCDSSTVTLRNGIESMLMHYIEEVQGVEQVLD 291 H.sapiens QEDGGDVIYKGFED--GIVQLKLQGSCTSCPSSIITLKNGIQNMLQFYIPEVEGVEQVMD 243 S.cerevisiae LEDGGDIDYRGWDPKTGTVYLRLQGACTSCSSSEVTLKYGIESMLKHYVDEVKEVIQIMD 229 R.prowazekii TQDGGDIIYKGFES--GVVKLALRGACLGCPSSTITLKNGIESMLKHFIPEVQEVKAVEE 187 :****: ::*:: * * * *:*:* * ** :**: **:.** .:: **: * : :

A.nidulans EEEEISMLEFAKFEEKLRQQKGAQATAPSSLDSAP 326 H.sapiens DE------SDEKEANSP------254 S.cerevisiae PEQEIALKEFDKLEKKLESSKNTSHEK------256 R.prowazekii DF------K------190

B NIFU Domain 1 Domain 2 Domain 3

TIND Domain 3

C An 220 MIKELLETRIRPAIQEDGGDIEFRGFENGIVMLKLRGACRTCDSSTVTLRNGIESMLMHYIEEVQGVEQV 289 I L RP DGGD E IV L GAC C SST TL IES L I VE V Aa 230 LIQKVLDEEVRPVLIADGGDVELYDVDGDIVKVVLQGACGSCSSSTATLKIAIESRLRDRINPSLVVEAV 300 * *

Figure 3.8. TIND is a highly conserved protein. (A) TIND homologues are present in organisms from bacteria through to humans. A ClustaW protein alignment of TIND homologues is shown. (B) TIND contains the carboxy- terminal domain of NIFU. (C) A ClustalW protein alignment between the carboxy-terminal domain of TIND (An) and the carboxy-terminal domain of NIFU from the nitrogen fixing bacteria Anabaena azollae (Aa) is shown. The * * indicate the location of two conserved cysteine residues which are present in the catalytic site of thioredoxin. 114

Figure 3.9. TIND localization. Cells which expressed TIND-HA were germinated in minimal media glycerol, fixed and processed for immunofluorescence. (A) TIND localization was examined using α-HA antibodies (3F10) followed by AlexaFluor 488 conjugated anti-rat IgG (Molecular

Probes). (B) To examine mitochondria, cells were incubated in minimal media containing 25nM MitoTracker (Molecular Probes) for 30 minutes prior to fixation.

(C and D) Cells expressing TIND GFP were germinated in minimal media, fixed, and processed for immunofluorescence. DNA was stained with DAPI and mitochondria were visualized by incubation in minimal media with 25nM

MitoTracker (Molecular Probes) for 30 minutes. (E) A. nidulans transformants which expressed TIND-GFP were germinated in minimal media. TIND-GFP localization was examined in living cells using a spinning disk confocal microscopy system (Perkin Elmer).

115 A α-HA (TIND)

B Mitotracker

C TIND-GFP D TIND-GFP

MitoTracker MitoTracker

DAPI DAPI

E

116 A TIND-GFP B

Tubulin

DAPI

C

Figure 3.10. TIND is mitochondrial during interphase and mitosis. To

determine TIND localization through the cell cycle. Cells which expressed TIND-

GFP were germinated in minimal media, fixed, and processed for

immunofluorescence. DNA was stained with DAPI. Microtubules were

visualized with mouse monoclonal antibody TAT-1 followed by AlexaFluor 594

conjugated rat anti-mouse IgG (Molecular Probes). Cells were determined to be

in (A) interphase, (B) metaphase, or (C) telophase based on DNA condensation and microtubule morphology.

117 1st M 2nd M 3rd M A ATG AAG ATG

B ATG ATG ATG

63bp insert w/ STOP

C ATG ATG ATG

Figure 3.11. Isolation of human tinD cDNAs. 5’ RACE PCR conducted in a

HeLa cDNA library identified three groups of human TIND cDNAs. (A) The most prevalent cDNA contained the first ATG but lacked the second. (B) Another class of cDNA contained all three in frame ATGs. (C) A third class, contained all three ATGs but a 63 bp insertion between the first and second ATGs contained an in frame STOP, suggesting that translation may initiate from the second ATG.

118

CHAPTER 4

TINC / An-HETC FUNCTION IS NON-ESSENTIAL

4.1. Introduction

The lack of multi-allelism at either the tinC of An-hetC loci suggests that neither of these genes function in heterokaryon incompatibility in A. nidulans. In order to examine the consequences of a loss of tinC function in A. nidulans, tinC was deleted. The conservation of tinC and An-hetC homologues in filamentous fungi suggests that they fulfill an important function in these organisms. The presence of the highly related An-hetC suggested a potential redundancy of function between these two loci, so both tinC and An-hetC-were deleted singly and in combination. These strains were examined for phenotypes under an array of growth conditions and different genetic backgrounds. Since a new technique was used to generate the deletion constructs, I will give a brief overview of the procedure.

4.1.1. A. nidulans transformation characteristics.

Transformation of plasmids and linear DNA in A. nidulans results in the integration of the transforming DNA into the genome of the recipient cell. The

119 degree of site specific homologous recombination of the transforming DNA can

be enhanced by plasmid linearization and by increasing the regions of

homologous DNA in the transformed DNA (Bird and Bradshaw, 1997). Typically

2 kB of homologous flanking sequence is deemed appropriate for high frequency

homologous targeting. While the recent availability of the A. nidulans genome

sequence has made restriction mapping of flanking regions more straightforward,

the generation of deletion cassettes was still cumbersome.

4.1.2. A method for generating deletion targeting constructs

Several studies have emphasized the utility of bacterial strains containing

inducible recombination machinery to generate recombinant plasmids or cosmids

for the purposes of gene deletion and gene tagging (Chaveroche et al., 2000; Yu

et al., 2000; Swaminathan et al., 2001) The E. coli strains, DY380 and DY331

were used in this study, contain temperature inducible recombination genes exo, bet, and gam (Yu et al., 2000). This strain displays very high recombination

efficiencies such that only 50 bp of homology is sufficient for homologous

recombination. In brief a deletion cassette is generated by PCR using primers

containing 50 bp of homology to sequences flanking the targeted locus followed

by 20 bp of homology to opposite termini of a bifunctional deletion cassette. The

availability of pyrG (Yu et al., 2000) and pyroA (Dou et al., 2003) (both genes

complement auxotrophic mutations in A. nidulans) deletion cassettes makes the

use of this method practical in A. nidulans. A plasmid or cosmid containing the

gene of interest is transformed into these recombinegenic E. coli strains. Cells

120 containing the plasmid or cosmid of interest were induced for recombination and

the deletion cassette is transformed into the strain. Following homologous

recombination of the deletion cassette into the genomic construct, the cosmid or

plasmid can be recovered and transformed into A. nidulans.

Note, at the time these studies were performed a 13X coverage of the A.

nidulans genome was not available. The current availability of a complete

genome sequence in combination with a Three Way Fusion PCR protocol has

improved on the plasmid recombination method described here. This method

has been used to great effect in gene targeted tagging and deletion studies

performed in our lab (Yang et al., 2004 Submitted).

4.2. Results

4.2.1. Generation of ∆tinC, ∆An-hetC, ∆tinC/∆An-hetC strains

tinC was deleted in A. nidulans using the method described above.

Endogenous tinC was successfully replaced with a pyrG / zeoR cassette. PCR was used to initially confirm which strains were tinC deleted. Primer set AO63 and AO74 amplified a 421 bp fragment if endogenos tinC was present. Primer set AO158 and AO165 amplified a 540 bp fragment if the deletion construct was present. Strains in which a 540 bp fragments was amplified with no 421 bp fragment were regarded as potential tinC deleted strains. Figure 4.1A shows

PCR analysis of nine potentially deleted strains and and R153 control strain.

Three of these nine strains contained a deletion specific PCR band and lacked a

121 tinC specific PCR fragment (Figure 4.1A). Figure 4.1B depicts an α-TINC antibody western blot of five potentially deleted strains. Note that while strains 2,

12, 13, and 20 lack TINC, strain 11 still contains TINC (Figure 4.1B). This result

is in agreement with the earlier PCR screen of these strains. The ability to

generate tinC deleted strains indicates that tinC function is unessential.

Deletion of An-hetC was achieved in generally the same manner as described for tinC. Importantly An-hetC was replaced with a zeoR/pyroA cassette

rather than the zeoR/pyrG cassette used for tinC. Lack of An-hetC was confirmed by PCR analysis and southern blotting (Data not shown). The ability to generate

An-hetC deleted strains indicates that An-hetC function is unessential.

Strains deleted for both tinC and An-hetC were obtained by crossing a tinC deleted strain with an An-hetC deleted strain. As described for tinC and An- hetC, the absence of TINC was confirmed in these strains by western blotting with α-TINC antbodies (Figure 4.1C). Lack of An-hetC was confirmed by PCR analysis (Figure 4.1C).

4.2.2. Deletion phenotype testing

∆tinC, ∆An-hetC, ∆tinC/∆An-hetC strains were examined under a range of conditions to determine whether loss of tinC/An-hetC function would generate any discernable phenotypes. These strains were able to undergo sexual crosses and produced viable meiotic progeny. 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

122 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. Lack of tinC did produce osmotic sensitivity, as ∆tinC strains were inhibited on YAGUU 1M sucrose (Figure 4.2A) or YAGUU 1M sodium chloride (Figure 4.2B). ∆tinC strains were also cold sensitive as they grew poorly at 20°C (Figure 4.2C). Due to this fact, ∆tinC cells were germinated in liquid media at 20°C and germinated in YG including 1M sucrose and examined by DAPI staining and tubulin immunofluorescence for potential mitotic defects. No significant defects were observed in any of these strains. Also, experiments described later in this thesis demonstrate that ∆N-TINC expression produces a striking defect in nuclear envelope severing. To examine the nuclear envelope in ∆tinc/∆An-hetC cells, this strain was crossed to a strain which expresses a GFP fusion protein with the nucleoporin SONAGle2/RAE1. Examination of nuclear envelopes in this strain revealed no defects in nuclear envelope severing

To further test for a cell cycle specific function for tinC strains were generated by mitotic crossing between ∆tinC/∆An-hetC and a number of A. nidulans cell cycle mutants Strains JD124 and JD127 were crossed to a number of strains carrying the following mutations: nimA1, nimA5, nimA7, nimX1 (CDK1), nimX2, nimX3, nudA5 (cytoplasmic dynein), nudA7, nimT23 (Cdc25), and bimE7

(APC1). These triple 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). No significant synthetic lethalities were detected.

123 4.3. Discussion

The fact that loss of tinC and An-hetC was not lethal in A. nidulans was

somewhat surprising given the presence of two related proteins in A. nidulans

and the apparent conservation of both an An-hetC and tinC homologue in filamentous fungi. Particularly disappointing was the lack of any observed cell cycle defects in any of the deletion strains even under environmentally stressful conditions. The inability to generate cell cycle specific defects in these null mutants leaves open the question as to whether ∆N-TINC represents a dominant negative form of TINC. While the deletion results argue against this idea they do not rule it out. For example, TINC may fulfill its function as part of a multi-protein complex. While loss of tinC can be effectively complemented by other complex members, the presence of ∆N-TINC in this complex may disrupt its function.

Alternately, tinC and An-hetC may fulfill an essential function under conditions not examined in these studies. Finally, additional tinC/An-hetC functional homologues with little primary sequence similarity may exist in A. nidulans.

The fact that ∆tinC growth is inhibited on 1M sodium chloride or 1M sucrose containing media suggests a potential role for TINC in osmoregulation in

A. nidulans. However, the fact that ∆tinC strains are also inhibited at 20°C suggests that these conditions may reflect general stress conditions. Under this premise, strains which lack tinC are compromised but these defects are masked under optimal growth conditions. Exposure of ∆tinC to general stress conditions including high osmolarity and cold effectively unmasks underlying defects.

124 In order to further investigate the role of TINC in A. nidulans, a synthetic lethality screen using ∆tinC and ∆tinC / ∆An-hetC should prove useful. The characterization of these genetic interactors should provide insight into the role of

TINC. A further discussion of potential roles for TINC based on additional experimental evidence is presented later in this thesis.

125 3 5 1 A R #11 #12 #13 #14 #15 #16 #17 #18 #20

tinC deleted wt tinC

1 1 2 3 0 B 2 # 1 1 2 # # # # C C N C C C 3 I N N N IN I I I 5 T 1 T t T T T ∆ ∆ ∆ ∆ R w TINC

α-tubulin

3 TINC Western Blot C 5 1 R 1 23465 798 10

TINC

C C A B An-hetC PCR BA t w ∆ An-hetC deleted

wt hetC

Figure 4.1 Deletion of ∆tinC and ∆An-hetC. (A) PCR screen of potentiall tinC deleted strains. Strain R153 was used as a wild type control. (B) Western blot of potentially tinC deleted strains probed with α-TINC antibodies. α-Tubulin was used as a loading control. (C) Confirmation of loss of tinC and An-hetC in double deleted strains.

126 GR5 R153 A

wt ∆tinC tinC wt ∆hetC hetC ∆tinC ∆tinC ∆hetC ∆hetC ∆tinC ∆tinC ∆hetC ∆hetC

B C

Figure 4.2. ∆tinC cells are osmotically sensitive and cold sensitive. (A)

∆tinC strains were inhibited on media containing 1M sucrose. (B) ∆tinC strains were inhibited on media containing 1M sodium chloride. (C) ∆tinC strains were inhibited at 20°C.

127

CHAPTER 5

TINC, TIND, AND NUCLEAR MEMBRANE FISSION

5.1. Introduction

Two-hybrid NIMA interacting cDNAs were secondarily screened by

overexpression in A. nidulans. Overexpression of NIMA induces premature

mitosis while a lack of functional NIMA results in an arrest in G2. Given this information it was reasoned that expression of NIMA interacting proteins which increased NIMA activity would promote premature mitosis while those which inhibited NIMA function would result in a nim defect. While these possibilities do not cover the array of potential NIMA interacting proteins it was reasoned that this would provide a good first step to indicate which of the NIMA interacting proteins were potentially involved in cell cycle regulation in A. nidulans. I have demonstrated previously that overexpression of ∆N-TINC and ∆N-TIND in A. nidulans produced a cell cycle specific defect characterized by polarized growth of the germling with an apparent lack of nuclear division. This defect in large part appeared identical to a nim phenotype in A. nidulans which can result from a lack of functional NIMA. Cells lacking functional NIMA undergo polarized growth and

128 arrest in late G2 prior to mitotic entry. These nuclei contain duplicated uncondensed DNA and visible nucleoli.

This chapter describes the further characterization of these phenotypes through the use of DAPI staining and tubulin immunofluorescence, as well as the use of a nucleoporin-GFP fusion protein, GFP-SONBNup98, as a new marker for cell cycle progression and nuclear envelope. Use of this fusion protein revealed that expression of ∆N-TINC or ∆N-TIND produced striking defects in nuclear envelope severing following mitosis, a process referred to as nuclear membrane fission. I will provide a brief introduction concerning the nature of the closed mitosis of A. nidulans and its consequences for cytokinesis and nuclear membrane fission, as well as, a description of the characteristics of the GFP-

SONBNup98 fusion protein and its utility in A. nidulans.

5.1.1. Closed mitosis

In vertebrate cells, one of the central hallmarks of mitosis is the breakdown of the nuclear envelope at prometaphase to allow for the interaction between chromosomes with cytoplasmic regulators including capture of kinetochores by mitotic spindle microtubules. The phosphorylation of lamins which exist as a meshwork of intermediate filaments underlying the nuclear

envelope is considered to be one of the primary triggers for nuclear envelope

breakdown (Marshall and Wilson, 1997). Recent studies have also implicated

cytoplasmic dynein in the process of physically tearing the nuclear envelope

apart (Beaudouin et al., 2002; Gonczy, 2002; Salina et al., 2002). During late

129 anaphase/telophase the nuclear envelope is reestablished through a stepwise

process involving association of nucleoporins and membrane components

around decondensing chromosomes. Work by several groups has indicated a

role for importins, nucleoporins and Ran GTPase in coordinating reformation of a

functional nuclear envelope following mitosis (Askjaer et al., 2002; Boehmer et al.,

2003; Harel et al., 2003a; Harel et al., 2003b; Ryan et al., 2003; Walther et al.,

2003a; Walther et al., 2003b; Wozniak and Clarke, 2003; Hachet et al., 2004). In vertebrate cells the combination of spatial separation of sister chromatids by the mitotic spindle coupled with the subsequent reformation of independent nuclear envelopes around these two separate masses of DNA results in the formation of two distinct daughter nuclei.

In contrast to the open mitosis exhibited by vertebrate cells, many fungi, including A. nidulans, undergo a closed mitosis, in which the nuclear envelope remains intact throughout the cell cycle (Robinow and Caten, 1969). In the absence of nuclear envelope breakdown, the mitotic machinery and regulators must be able to traverse this barrier to gain access to chromosomes and establish the mitotic spindle within the nucleus. Studies in A. nidulans have

identified the NIMA kinase as a key regulator of nuclear pore structure and

permeability at mitosis (Wu et al., 1998; De Souza et al., 2003; De Souza et al.,

2004 Submitted). Additionally, the lack of nuclear envelope disassembly and

reassembly requires that organisms which undergo a closed mitosis be able to

coordinate nuclear envelope severing, or nuclear membrane fission, with mitotic

exit. (Latterich et al., 1995)

130 5.1.2. Nuclear membrane fission

As described previously, vertebrate cells undergo nuclear breakdown at

mitotic onset and the nuclear envelope reforms around separated DNA during

mitotic exit (Figure 5.1 Humans). This cycle of nuclear envelope breakdown and

reformation negates the requirement for nuclear membrane fission

The lack of nuclear envelope disassembly and reassembly requires that

organisms undergoing a closed mitosis be able to coordinate nuclear envelope

severing, or nuclear membrane fission, with mitotic exit. The mechanisms

underlying this process are not well understood. It has been suggested that

nuclear membrane fission may proceed in a manner analogous to the Cdc48

directed homotypic membrane fusion and fission of the endoplasmic reticulum

(Latterich et al., 1995). Studies in S. cerevisiae examined the role of cytokinesis

in driving nuclear membrane fission. The authors of this study examined nuclear

envelope morphology in mutants defective in cytokinesis. These experiments

demonstrated that 70% of the mutants initiated a second round of budding in the absence of nuclear membrane fission. DNA in the mother cell and bud were contained within the same nuclear envelope (Lippincott and Li, 2000). These experiments suggest a role for cytokinesis in driving nuclear membrane fission

(Lippincott and Li, 2000) (Figure 5.1 S. cerevisiae). In S. cerevisiae every mitotic division is followed by cytokinesis (Figure 5.1 S. cerevisiae). This is not the case

in A. nidulans where, in order to produce a multi-nucleate cell, rounds of mitotic

division (with nuclear membrane fission) can occur in the absence of cytokinesis

or septation (Fiddy and Trinci, 1976; Harris et al., 1994; Wolkow et al., 1996;

131 Harris et al., 1997) (Figure 5.1 A. nidulans). During these divisions nuclear

membrane fission must be mediated through mechanisms not linked to

cytokinesis (Figure 5.1 A. nidulans).

5.1.3. Examination of the nuclear envelope in A. nidulans

Previous work in the lab has identified two components of the nuclear pore

complex (NPC), SONAGle2/RAE1 and SONBNup98 as NIMA interacting proteins (Wu et al., 1998; De Souza et al., 2003). The NPC is a large macromolecular protein complex which spans the nuclear envelope and allows transport of proteins and nucleic acids between the nucleus and the cytoplasm (Wente, 2000; Rout and

Aitchison, 2001). Nup98 is an FG repeat protein which plays a role in regulating transport through the nuclear pore. To examine the localization of SONBNup98, Dr.

Colin De Souza generated a strain which expresses a GFP-SONBNup98 fusion

protein from the endogenous sonB promoter (CDS165). Examination of this

strain using a confocal live cell imaging system revealed that, as expected GFP-

SONBNup98 appeared in fluorescent rings throughout the length of the germ tube with spacing and sizes expected for nuclei. Additionally, a small percentage of these cells displayed a diffuse cytoplasmic fluorescence with no concentration at the nuclear periphery. Live cell imaging of these cells revealed that GFP-

SONBNup98 was present at the nuclear periphery during the vast majority of the

cell cycle but that it was strikingly lost at mitosis. At mitosis GFP-SONBNup98 was observed throughout the cytoplasm and within nuclei. Following mitosis GFP-

SONBNup98 reassociated with nuclear envelopes. GFP-SONBNup98 localization

132 was confirmed in fixed cells which allowed for DNA staining and examination of

spindle morphology to determine cell cycle stage. These studies confirmed that

GFP-SONBNup98 was present around DNA until metaphase and that GFP-

SONBNup98 reappeared at the nuclear periphery during telophase.

5.2. Results

5.2.1. ∆N-TINC expression but not TINC expression produces nuclear

division defects

As shown previously, expression of the 5’ truncated tinC cDNA obtained

from the Two-hybrid screen produced growth inhibition with an underlying cell

cycle defect in A. nidulans. In order to confirm this defect and to test whether

expression of full-length TINC caused a similar defect, genomic clones

corresponding to the 5’ truncated cDNA already isolate and a full-length genomic

tinC clone were amplified by PCR and cloned into pAL3. These constructs were

transformed into A. nidulans and transformants randomly selected and tested on inducing and repressing media (Figure 5.2). As expected control transformants containing only empty pAL3 vector grew generally well on both inducing and repressing media (Figure 5.2 alcA::). Also, expression of a genomic 5’ truncated tinC clone produced the variable growth inhibition seen during expression of 5’ truncated tinC cDNA (Figure 5.2 ∆N-TINC). Disappointingly, expression of full- length TINC failed to produce significant growth inhibition (Figure 5.2 TINC). The

133 few colonies which were repressed on inducing media also grew poorly on

repressing media.

Cells expressing ∆N-TINC were germinated on coverslips in minimal

media with ethanol at 30°C and DAPI stained to examine nuclear morphology.

As described previously, empty vector control cells displayed nuclear morphology

and division rates equivalent to wild-type controls (Figure 5.3A a). Also as

expected expression of ∆N-TINC caused nuclear division defects (Figure 5.3A b-

f). A closer examination of these cells revealed that nuclei of several different nuclear phenotypes existed. Significantly, 68% of ∆N-TINC-HA germlings exhibited a nim phenotype (Figure 5.3A b and Figure 5.3B Single), typical of cells which lack functional NIMA. These cells displayed polarized growth in the absence of nuclear division. Typically these nuclei were large with uncondensed

DNA and a visible nucleolus, all characteristics of late interphase nuclei (Figure

5.3A b). The extremely large size of some of these nuclei suggested that they had undergone multiple rounds of DNA replication. The nuclear morphology of the approximately 32% remaining germlings also suggested that these cells had not merely arrested in late interphase (Figure 5.3A c-f and Figure 5.3B). 3% of cells examined contained condensed DNA (Figure 5.3A c and Figure 5.3B).

Another class of mutants contained two discernable DNA masses, although these nuclei frequently appeared abnormal. Specifically, nuclei were frequently of uneven sizes or appeared stretched (Figure 5.3A d and Figure 5.3B Separated

Unlinked). In 11% of the total germlings separated masses of DNA were linked by intervening DNA (Figure 5.3A e and Figure 5.3B Separated Linked). Finally,

134 8% of ∆N-TINC-HA germlings displayed fragmented DNA (Figure 5.3A f and

Figure 5.3B Fragmented).

5.2.2. ∆N-TINC is present in membranes

In order to examine the subcellular localization of ∆N-TINC, a strain which expressed ∆N-TINC-HA (JD155) under the control of the alcA promoter was germinated on coverslips in inducing media. Germlings were fixed and processed for immunoflouresence using anti-HA antibodies (3F10), tubulin antibodies (TAT1), and DAPI staining. ∆N-TINC was present in a punctate pattern throughout the cytoplasm punctuated by concentrations of ∆N-TINC at the periphery of circular structures, presumably membranes (Figure 5.4A ∆N-

TINC). No anti-HA signal was detected in a control strain containing empty vector(JD148) (Figure. 5.4A control).

An additional strain expressing higher levels of ∆N-TINC-HA (CB45) was also examined for ∆N-TINC-HA localization. This strain was inoculated on coverslips in minimal media glucose and allowed to germinate. After germination cells were shifted to inducing media and time course samples removed for immunofluorescence at one, three, and six hours following induction. Following one hour of induction ∆N-TINC was observed in a faint punctate pattern throughout the cytoplasm (Data not shown). After three hours of induction ∆N-

TINC was visible in a punctate pattern in the cell and concentrated at the periphery of circular membranes throughout the length of the cells (Figure 5.4B

+3 hours). Following six hours of induction, the membranes marked by ∆N-TINC

135 localization appeared to have coalesced into irregular membrane aggregates

which were associated with nuclei in these cells (Fig. 5.4B +6 hours).

In an attempt to observe the formation of these membrane aggregates a

construct was created which produced ∆N-TINC-GFP under the control of the

alcA promoter. Examination of ∆N-TINC-GFP in transformed strains germinated in inducing media demonstrated that ∆N-TINC-GFP was present inside rather than at the periphery of membranes (Figure 5.4C). Also, expression of ∆N-TINC-

GFP failed to produce the colony growth defects and underlying cell cycle defects seen for ∆N-TINC expression.

5.2.3. ∆N-TIND expression but not TIND expression produces nuclear

division defects and abnormal mitotic spindles

As shown previously, expression of the 5’ truncated tinD cDNA obtained from the Two-hybrid screen produced growth inhibition with an underlying cell cycle defect in A. nidulans. In order to confirm this defect and to test whether expression of full-length TIND caused a similar defect, genomic clones for corresponding to the 5’ truncated cDNA already isolate and a full-length genomic tinD clone were amplified by PCR and cloned into pAL3. These constructs were transformed into A. nidulans and transformants randomly selected and tested on inducing and repressing media (Figure 5.5). As expected control transformants containing only empty pAL3 vector grew well on both inducing and repressing media (Figure 5.5 alcA::). Also expression of a genomic 5’ truncated tinD clone produced the variable growth inhibition seen for expression of 5’ truncated tinD

136 cDNA (Figure 5.5 ∆N-TIND). Also, expression of full-length TIND produced significant growth inhibition (Figure 5.5 TIND).

Cells expressing ∆N-TIND and TIND were germinated on coverslips in minimal media with ethanol at 30°C and DAPI stained to examine nuclear morphology. As described previously, empty vector control cells (JD157) displayed nuclear morphology and division rates equivalent to wild-type controls

(Figure 5.6A a). Cells expressing TIND displayed very poor spore viability with the vast majority of cells failing to germinate (Data not shown). Examination of other strains expressing lower levels of TIND revealed that while these cells grew extremely slowly, they did undergo nuclear division (Data not shown).

Examination of cells expressing ∆N-TIND (JD159) displayed a mix of mutant phenotypes strikingly similar to those seen for cells expressing ∆N-TINC. 57% of these cells displayed a single DNA mass, which as described for ∆N-TINC were typically large and contained uncondensed DNA (Figure 5.6A b and Figure 5.6B

Single). Additionally, 3% of these cells contained a single mass of condensed

DNA. 10% of cells contained two or more masses of DNA with visible DNA linkage (Figure 5.6A d and Figure 5.6B Separated Linked). 29% of cells contained two or more masses of DNA with no discernable linkage (Figure 5.6A c and Figure 5.6B Separated Unlinked). These masses of DNA were frequently of unequal size and of abnormal number. The cell pictured contains three apparent nuclei which would not exist in a wild type cell, as cells proceed from two nuclei to four nuclei following mitosis (Figure 5.6A c). Finally, 4% of cells contained fragmented DNA (Figure 5.6A e and Figure 5.6B Fragmented).

137 In addition to nuclear morphology defects, expression of ∆N-TIND produced defects in mitotic spindle morphology. In control cells metaphase spindles appeared as bars with a mass of condensed DNA present between the spindle poles (Figure 5.7A). Spindle morphology was examined in two strains of cells which express either ∆N-TIND (JD73) (Figure 5.7B and C) or ∆N-TIND and tubA-GFP (JD ) (Figure 5.7D-G). Cells which expressed ∆N-TIND contained defective mitotic spindles which appeared similar to monopolar spindles (Figure

5.7B-G). These spindles appeared either in a “V-shaped” configuration (Figure

5.7B-F) or as a single bar which was ill-defined at one end (Figure 5.7G and H).

DNA frequently extended beyond the limits of the spindle.

5.2.4. ∆N-TIND is present throughout the cell

A transformant which expressed ∆N-TIND-HA (JD73) under the control of the alcA promoter was germinated under inducing conditions in liquid media.

After allowing cells to grow to germling size, they were fixed, and ∆N-TIND-HA localization examined using an anti-HA antibody. HA epitopes were detected throughout the entire cell both in the cytoplasm and the nuclei (Figure 5.8A and

B).

5.2.5. Cells expressing ∆N-TINC or ∆N-TIND proceed through faulty mitoses

In order to further characterize the cell cycle defects produced by expression of ∆N-TINC and ∆N-TIND, alcA::∆5’tinC-HA and alcA::∆5’tinD-HA constructs were transformed into A. nidulans strain CDS165. Strain CDS165

138 expresses a GFP fusion protein with the nucleoporin, SONBNUP98 which allows for visualization of the nuclear envelope through much of the cell cycle (De

Souza et al., 2003). GFP-SONBNUP98 also acts as a marker of cell cycle progression since GFP-SONBNUP98 is present at the nuclear envelop through

interphase but is dispersed during mitosis (De Souza et al., 2003).

As previously described, transformants were screened on inducing media

to identify strains which expressed ∆N-TINC or ∆N-TIND at levels high enough to

produce cell cycle defects. Empty vector control cells (JD157) and strains

expressing either ∆N-TINC (JD158) or ∆N-TIND (JD159) were germinated at

30°C in mmEthanol on coverslips, fixed and stained with DAPI and processed for immunofluorescence using an α-tubulin antibody to confirm that ∆N-TINC-HA expressing cells were attempting to undergo mitosis.

As expected, approximately 95% of empty vector control cells (JD157)

displayed interphase cytoplasmic microtubules and nuclei with GFP-SONBNUP98 at their periphery (Figure 5.9A). GFP-SONBNUP98 was dispersed in control cells

at metaphase (Figure 5.9B) and reassociated at telophase (Figure 5.9C). Cells

expressing either ∆N-TINC or ∆N-TIND also displayed GFP-SONBNUP98 fluorescence around the periphery of interphase nuclei (Figure 5.9D and G).

Additionally, mitotic cells had dispersed GFP-SONBNUP98 throughout the cell

(Figure 5.9E, F, and H). Spindles in ∆N-TINC expressing cells looked similar to

wild-type telophase spindles. The spindles were typically stretched and thin

(Figure 5.9E). Dense bundles of astral microtubules were also present frequently

(Figure 5.9F). Cells expressing ∆N-TIND contained defective spindles as

139 described previously. Note that the abnormal spindle and DNA morphologies

seen in ∆N-TIND and ∆N-TINC expressing cells made it difficult to determine

how far cells had progressed through mitosis, however, the ability to generate

spindles, undergo chromosome condensation and disperse GFP-SONBNUP98 during mitosis confirmed that these cultures are progressing through defective mitoses.

5.2.6. Cells expressing ∆N-TINC or ∆N-TIND display nuclear membrane

fission defects

Expression of ∆N-TINC and ∆N-TIND in CDS165 also provided the opportunity to examine the nuclear envelope in cells which by DAPI staining appear to contain two discreet masses of DNA. The presence of “linking” DNA between separated DNA masses in cells which expressed ∆N-TINC or ∆N-TIND, suggested that these cells were not undergoing nuclear division properly. During closed mitoses, the production of two independent nuclei is accomplished though cleavage of the nuclear envelope. The presence of intervening DNA suggested that cells expressing ∆N-TINC or ∆N-TIND may be defective for nuclear membrane fission.

To explore this possibility, the nuclear envelopes in these cells were

studied by examining GFP-SONBNUP98 localization in control cells or cells which expressed ∆N-TINC or ∆N-TIND. Cells were identified, which by DAPI staining contained two or more DNA masses. In some cases, DNA was visible trailing between the nuclei, but in some cases it was not. GFP-SONBNUP98 localization

140 was examined in these cells to determine whether, apparently separate DNA

masses were contained within a single nuclear envelope. Cells classed as

containing “Normal nuclear envelopes” contained a single DNA mass within each

nuclear envelope (defined by GFP-SONBNUP98 localization). Alternatively, cells in which two or more masses of DNA were present within a single nuclear envelope were classified as containing “Stretched nuclear envelopes”.

For control cells, each separate DNA mass was contained within a nuclear envelope (Fig. 5.10A vector and C, Normal nuclear envelope). However, in cells which expressed ∆N-TINC, nuclear membrane fission was defective. Specifically,

71% of cells which by DAPI staining contained separated DNA masses had failed to undergo nuclear membrane fission (Fig. 5.10A ∆N-TINC and C, Stretched nuclear envelope). In these cells, two or more DNA masses were contained within a single nuclear envelope. Cells that expressed ∆N-TIND displayed a similar defect in nuclear membrane fission as 66% of cells which by DAPI staining contained separated DNA masses had failed to undergo nuclear membrane fission (Fig. 5.10A ∆N-TIND and C, Stretched nuclear envelope).

In some cases this nuclear envelope linkage between DNA masses was maintained over long distances (Figure 5.10B). Figure 5.10B is an image of

GFP-SONBNUP98 localization in a living cell that expressed ∆N-TINC. The arrows indicate bulges within the nuclear membrane which presumably correspond to

DNA masses. In this image four DNA masses are present within a single nuclear envelope which extends ~40 µm.

141 5.3. Discussion

5.3.1. TINC, TIND and growth inhibition

Initial expression of tinC and tinD cDNAs recovered from the Two-hybrid screen produced variable colony growth inhibition when expressed in A. nidulans.

The initial cloning strategy for the Two-hybrid cDNAs resulted in the addition of eight amino acids homologous to the carboxy terminus of the GAL4-AD downstream from an artificial start of translation. To ensure that the overexpression defects were not due to the presence of these extra amino acids, similarly truncated genomic clones lacking these GAL4-AD amino acids were expressed in A. nidulans. These genomic constructs showed the same growth

defect observed in earlier experiments. Disappointingly, however, expression of

full-length TINC failed to produce growth inhibition in A. nidulans demonstrating

that the generation of these defects is restricted to truncated forms of TINC. This

result suggests that ∆N-TINC may represent a dominant negative form of TINC,

a possibility that will be discussed later. In contrast to TINC, the expression of

TIND was quite lethal, prompting further examination of these cells to determine

if the underlying defect was cell cycle specific.

5.3.2. Cell cycle specific defects

While the vast majority of nuclei in ∆N-TINC and ∆N-TIND expressing

cells resembled late interphase nuclei, the characteristics of some of the nuclei in

these cells, including large size, DNA which appeared condensed, and highly

142 stretched nuclei, suggested that they were not merely arresting in G2. These additional phenotypes prompted a more detailed characterization of the ∆N-TINC and ∆N-TIND defects. Close examination of the DAPI staining allowed for the classification of individual germlings based on their nuclear morphology. The four types were: 1) single nucleus, 2) two or more separated nuclei, 3) separated nuclei with intervening DNA, or 4) fragmented DNA. The morphology of some of these nuclei suggested that these cells were progressing through an aberrant cell cycle rather than arresting in late interphase. First, while the majority of cells did contain single large nuclei with uncondensed DNA in many cells the large size of nuclei suggested that the DNA had gone through DNA replication in the absence of mitotic division. Second, roughly 3-5% of the cells contained condensed DNA, suggesting that these cells were entering mitosis. The fact that the percentage of cells apparently in mitosis did not increase over time suggested that these cells are traversing mitosis rather than becoming arrested. Third, the fact that some cells contained two apparently independent masses of DNA suggested that these cells had completed nuclear division. It was noted, however, that these nuclei were frequently of uneven size and spacing suggesting underlying defects in chromosome separation and segregation. In addition to the nuclear division defects observed by DAPI staining, cells expressing ∆N-TIND contained spindle defects which were suggestive of monopolar spindles.

GFP-SONBNUP98 was used to great effect in these studies as both a nuclear envelope marker and to monitor cell cycle progression. Expression of

∆N-TINC and ∆N-TIND in a strain expressing GFP-SONBNUP98 provided further

143 evidence that these ells are progressing through mitosis as GFP-SONBNUP98 dispersal correlated with the presence of mitotic spindles. Note that while monopolar spindles were not observed in ∆N-TINC expressing cells, the spindles within these cells were aberrant with many of them appearing similar to telophase spindles in wild type cells. These long thin spindles may have been a byproduct of the large size of the nuclei. Some also appeared to contain multiple spindle pole bodies suggestive of these cells progressing through the cell cycle without nuclear division. Regardless, the ability of these cells to generate spindles and disperse GFP-SONBNUP98 suggests these cells are able to enter mitosis.

An attractive model for explaining these cell cycle defects is that ∆N-TINC and ∆N-TIND inhibit NIMA function. The fact that these are able to progress through mitosis suggests two things about this potential inhibition. First, while

TINC and TIND inhibit NIMA they do so to a level which still allows entry and progression through mitosis. Under this assumption the defects would be a result of progression through mitosis with inadequate NIMA. Second, alternatively, the inhibition could be specific to mitotic forms of NIMA. It is known that NIMA undergoes hyperphosphorylation to produce a highly active mitotic specific form of the kinase (Ye et al., 1995). TINC and TIND may selectively inhibit these forms of NIMA, allowing cells to enter mitosis but prematurely extinguishing NIMA activity

Examination of nuclear envelopes with GFP-SONBNUP98 was also informative in that examination of cells which contained apparently two or more

144 nuclei based on DAPI staining proved frequently to actually contain a single

nucleus based on the presence of GFP-SONBNUP98 both around the periphery of each DNA mass and interconnecting these masses of DNA. This result illustrates that in addition to spindle and chromosome segregation defects these cells are also defective in nuclear membrane fission. In wild type cells DNA is segregated on the mitotic spindle followed by cleavage of the nuclear envelope.

In these cells the DNA is segregated in some cases but the active process of nuclear envelope cleavage has failed.

Finally, although TIND produced severe growth inhibition, microscopic

examination revealed that these cells appeared to germinate very slowly if at all

under inducing conditions. The fact that the difference between TIND and ∆N-

TIND is the presence of a mitochondrial leader sequence in the former, suggests that TIND overexpression may overwhelm the mitochondrial transport capabilities and impair mitochondrial function. This idea is supported by the fact that expression of TIND to high levels results in TIND being localized throughout the cytoplasm.

5.3.3. ∆N-TINC and ∆N-TIND Localization

The expression of HA epitope tagged forms of ∆N-TIND and ∆N-TINC

was used to examine their localization in A. nidulans. While TIND was clearly

shown to be a mitochondrial protein, ∆N-TIND was observed throughout the cell.

This result provides good evidence that the predicted mitochondrial leader

sequence of TIND is indeed functional, since the amino-truncation point of ∆N-

145 TIND produces a form of TIND that lacks this leader sequence. This raises the

question why ∆N-TIND but not TIND induces cell cycle defects. One possibility is that in the absence of a mitochondrial leader sequence ∆N-TIND does not overwhelm the transport properties of the mitochondria and impair their function.

∆N-TINC was observed in a speckled pattern throughout the cytoplasm in

a pattern not unlike that seen for TINC. However, the hallmark of ∆N-TINC

localization was the accumulation of ∆N-TINC at the periphery of apparently

membranous structures. The examination of ∆N-TINC localization over time was

striking. One hour following induction ∆N-TINC was present throughout the cytoplasm with some larger foci. Over the next six hours these foci appeared to increase into spherical structures which apparently aggregated around nuclei.

The inability to observe the localization of ∆N-TINC over time in living cells was disappointing as it would have provided direct evidence for the “growth” of ∆N-

TINC-containing membranes rather than the transfer of ∆N-TINC between increasingly larger membranes. The expression conditions required to induce the formation of ∆N-TIND associated membranous aggregates resulted in the expression of ∆N-TIN-GFP to such a level that it was targeted to the interior of membranes which may represent vacuoles. Mislocalization of ∆N-TINC-GFP in these cells resulted in a failure to produce growth inhibition or cell cycle defects.

This mislocalization of GFP fusion proteins has been reported by others working in the lab (Dr. Colin De Souza personal communication).

It is important to note that the membrane aggregates observed in some cells are not required for the production of cell cycle defects. However, the

146 association of ∆N-TINC-containing structures with nuclei was intriguing, in that it suggested a possible connection between these membranes and the nuclear envelope. This fact prompted the examination of ∆N-TINC localization in a strain

which expressed GFP-SONBNUP98. While ∆N-TINC did not colocalize with the

GFP-SONBNUP98 this result does not rule out the possibility that these membranous ∆N-TINC containing structures are aberrant ER/nuclear envelope devoid of nucleoporins. Two previous studies have suggested a link between insufficient NIMA and nuclear envelope aberrations. First, A. nidulans cells which enter mitosis with inadequate NIMA display significant nuclear envelope perturbations (Osmani et al., 1991b). Second deletion of the S. pombe NIMA

related kinase, Fin1p results in generally abnormal nuclear envelope morphology

and excess nuclear envelopes.

Further interpretation of the nuclear membrane fission defects and their

potential connection to NIMA will be reserved for Chapter 7.

147 Mitotic G2 M Exit G1 Humans

Nuclear Nuclear envelope envelopes breakdown reform & cytokinesis

S. cerevisiae

cytokinesis & karyofission

A. nidulans

karyofission

Key

= cytokinesis = nuclear envelope

= DNA = cell membrane

Figure 5.1. Nuclear membrane fission. Nuclear membrane fission is the process of nuclear severing which occurs at the completion of mitosis in organisms which undergo a closed mitosis. No nuclear membrane fission occurs in human cells as the nuclear envelope breaks down at mitotic entry and reforms around separated chromosomes at mitotic exit. In S. cerevisiae nuclear membrane fission is thought to be driven in part by cytokinesis which occurs at the end of each mitosis. The absence of cytokinesis during the early rounds of nuclear division in A. nidulans necessitates an independent process of nuclear envelope severing at mitotic exit. 148 alcA:: ∆N-TINC TINC

mmGLU

mmEtOH

Figure 5.2. Expression of ∆N-TINC inhibits growth in A. nidulans. A. nidulans strains transformed with empty vector (alcA::), alcA::-5’tinC (∆N-TINC), or alcA::tinC (TINC) were spotted onto repressing (mmGLU) and inducing media

(mmEtOH).

149 A a DAPI b c

DIC

d e f

B 100

80 71%

60 Vector ∆N-TINC

entage of Cells 40 c

Per 20 11% 10% 8% 0 S S S F in e e r p U p a g L g le a n a m in r l ra k a in e e te k te n d d e d te d d

Figure 5.3. Expression of ∆N-TINC produces nuclear division defects. (A)

(a) Empty vector control cells and cells which expressed (b-f) ∆N-TINC were germinated in minimal media ethanol, fixed and stained, with DAPI. (a) Empty vector control cells displayed wildtype nuclear division. (b-f) Cells which expressed ∆N-TINC displayed a range of nuclear defects including: (b) a single large uncondensed nucleus, (c) a single nucleus with condensed DNA, (d) separated DNA, (e) separated DNA with visible DNA linkage, or (f) fragmented

DNA. (B) The percentage of cells displaying various nuclear morphologies was determined for the control and ∆N-TIND-HA expressing cells after DAPI staining.

150

Figure 5.4. ∆N-TINC locates to the periphery of membranes. (A) Cells which

expressed ∆N-TINC-HA (JD155) or control cells (JD148) were germinated in

inducing media, fixed and processed for immunofluorescence. ∆N-TINC-HA was visualized with anti-rat HA antibody 3F10 followed by mouse anti-rat AlexaFluor

488 IgG. Tubulin was visualized as described previously. Cells were DAPI stained to visualize DNA. (B) Cells which expressed ∆N-TINC-HA (CB45) were germinated under repressing conditions and then shifted to inducing media.

Following three (+3 hours) and six hours (+6 hours) of induction cells were fixed and processed for immunofluorescence. ∆N-TINC-HA was visualized as described in (A). Cells were DAPI stained to visualize DNA. (C) Cells which expressed ∆N-TINC-GFP were germinated in inducing conditions. ∆N-TINC-

GFP localization was examined in living cells using a spinning disk confocal microscopy system (Perkin Elmer).

151 A Control ∆N-TINC ∆N-TINC

Tubulin

DAPI

B +3 hours +6 hours DIC

∆N-TINC

DAPI

Merge

C

152 alcA:: NIMA TIND ∆N-TIND

mmGlu

mmEtOH

Figure 5.5. Expression of ∆N-TIND or TIND inhibits growth in A. nidulans. A. nidulans strains transformed with empty vector (alcA::), alcA::nimA (NIMA), alcA::-5’tinD (∆N-TIND), or alcA::tinD (TIND) were spotted onto repressing

(mmGLU) and inducing media (mmEtOH).

153 A a DAPI bc

DIC

de

B 100

80 s l Cel 60 57%

age of Vector

ent 40 ∆N-TIND c 29% Per 20 10% 10% 4% 0 S S S F in e e r p U p a g L a a g le in n r m ra li a k n t e e te k e n d d e d t d e d

Figure 5.6. Expression of ∆N-TIND produces nuclear division defects. (A)

(a) Empty vector control cells and cells which expressed (b-e) ∆N-TIND were germinated in minimal media ethanol, fixed and stained, with DAPI. (a) Empty vector control cells displayed wild type nuclear division. (b-e) Cells which expressed ∆N-TIND displayed a range of nuclear defects including: (b) a single large nucleus, (c) separated DNA, (d) separated DNA with visible DNA linkage or

(e) fragmented DNA. (B) The percentage of cells displaying various nuclear morphologies was determined for the control and ∆N-TIND-HA expressing cells after DAPI staining.

154 A B Tubulin C DIC

Tubulin DAPI

DAPI

Merge E DIC F

Tubulin

DAPI

Merge

G H

Figure 5.7. Expression of ∆N-TIND produces defective spindles. (A) Empty vector control cells and cells which expressed (B and C) ∆N-TIND (JD73) and cells which expressed (D-G) ∆N-TIND in a strain which also expressed tubulin-

GFP (JD166). Cells were germinated in minimal media ethanol, fixed and processed for immunofluorescence. DNA was stained with DAPI and tubulin was visualized with monoclonal TAT1 antibodies followed by Alexa Fluor 488 conjugated rat anti-mouse IgG (Molecular Probes) for strain JD73.

155 HA HA

DAPI DAPI

Figure 5.8. ∆N-TIND locates throughout the cell. Cells which expressed ∆N-

TIND-HA (JD73) were germinated in minimal media glycerol, fixed and processsed for immunofluoresence. Cells were stained with DAPI and ∆N-TIND localization was examined using anti-HA antibodies followed by AlexaFluor 488 conjugated rat anti-mouse IgG (Molecular Probes).

156 Interphase Mitosis A DAPI B C

or SONB-GFP Vect α-Tubulin

D E F TINC N- ∆

G H ND I N-T ∆

Figure 5.9. Cells expressing ∆N-TINC or ∆N-TIND proceed through faulty mitoses. Cells which expressed GFP-SONBNUP98 (JD157) (A, B, and C), GFP-

SONBNUP98 and ∆N-TINC-HA (JD158) (D, E, and F), or GFP-SONBNUP98 and ∆N-

TIND (JD159) (G and H) were germinated at 32°C in minimal media ethanol, fixed, and processed for immunofluorescence. Tubulin was visualized with TAT1 mouse monoclonal antibody followed by AlexaFluor 594 IgG. Interphase cells displayed uncondensed DNA, GFP-SONBNUP98 present around nuclei and interphase microtubules (A, D, and G). Mitotic cells contained condensed DNA, lacked GFP-SONBNUP98 (until telophase), and contained mitotic spindles (B, C, E,

F, G, and H).

157

Figure 5.10. Expression of ∆N-TINC or ∆N-TIND produces nuclear membrane fission defects. (A). Control cells (JD 157) (vector), ∆N-TINC expressing cells (JD158) (∆N-TINC), or ∆N-TIND expressing cells (JD159) (∆N-

TIND) were germinated in inducing media, fixed and stained with DAPI. GFP-

SONBNUP98 allowed visualization of nuclear membranes within the cells. (B).

Image of GFP-SONBNUP98 in a living cell expressing ∆N-TINC (JD158).

Illustrates that the nuclear envelopes in these cells can become dramatically stretched. Total length of nuclear envelope in this image ~40 µm. Arrows indicate the predicted positions of DNA masses. (C) Quantification of nuclear membrane fission defects in cells expressing ∆N-TINC or ∆N-TIND . GFP-

SONBNUP98 localization was examined in cells (vector, ∆N-TINC, and ∆N-TIND) which by DAPI staining appeared to contain separated DNA. Cells containing separated DNA were classed as one of the following based on GFP-SONB localization. First, cells in which each nuclear envelope contained a single DNA mass were said to contain “Normal nuclear envelopes”. Second, cells in which the two or more DNA masses were present within a single nuclear envelope were said to contain “Stretched nuclear envelope”.

158 A vector ∆N-TINC ∆N-TIND

DIC

DAPI

GFP-SONB

MERGE

B C 100

80 71% 66% 60

40 34% 29% Percentage of Cells vector 20 ∆N-TIND

∆N-TINC 0 S No e tr e n r n e m v tc v e h e a lo e lo l p d p n e e u n c u le c a le r a r

159

CHAPTER 6

TINC INTERACTS WITH NIMA IN A. NIDULANS

6.1. Introduction

We have identified TINC and TIND as NIMA interacting proteins based on their ability to bind NIMA in the yeast Two-hybrid screen. While these results, in combination with the cell cycle specific defects observed with TINC and TIND expression in A. nidulans, strongly suggest that TINC and TIND interact with

NIMA in A. nidulans, this must be confirmed by interaction studies in A. nidulans.

While the Two-hybrid screen does allow for the rapid identification of cDNAs which encode interacting proteins, it is not without drawbacks. First, the subcellular localization of GAL4 fusion proteins in the screen is often radically different from their endogenous localization. The addition of GAL4 to the amino- terminus of both bait and target proteins localizes theses fusion proteins to the nucleus. This effectively brings proteins in contact with each other which may normally be confined to completely separate subcellular compartments (example mitochondria or ER). Conversely, this mislocalization may produce the opposite effect, whereby two proteins which interact in an endogenous setting fail to interact in the Two-hybrid due to mislocalization in the nucleus where required

160 accessory factors required for interaction may not exist. Second, although A. nidulans and S. cerevisiae are both fungi, they are very different organisms. As a consequence of this fact, proteins or conditions required for protein interaction in A. nidulans may not be present in S. cerevisiae. Conversely, additional proteins or altered conditions in S. cerevisiae may produce protein interactions in the Two-hybrid which do not occur in A. nidulans. In the case of NIMA, it is not clear the degree to which NIMA is active in the yeast Two-hybrid screen, although the fact that recombinant NIMA produced in bacteria is active as a kinase suggests that NIMA is probably activated in yeast as well. Finally, the addition of GAL4 sequence to the amino-termini of proteins frequently results in a truncation of the amino terminus. The simple addition of this polypeptide coupled with potential truncations produces a protein which is potentially far different than the endogenous protein. Either of these circumstances could produce significant misfolding of the protein of interest.

The fact that the forms of TINC and TIND which interacted with NIMA in the Two-hybrid screen were amino-truncated, and that in both cases this truncation resulted in the loss of a putative amino-terminal targeting peptide, highlights the importance of demonstrating that these two proteins interact in A. nidulans. Studies described in this chapter demonstrate that while we were unable to confirm the TIND-NIMA interaction, TINC does interact with NIMA in A. nidulans. Additionally, it is shown that expression of ∆N-TINC apparently destabilizes NIMA in mitotic samples.

161 6.2. Results

6.2.1. TINC interacts with NIMA in A. nidulans

To confirm the interaction between NIMA and TINC we observed in the yeast Two-hybrid screen, we performed co-immunoprecipitation experiments from A. nidulans protein extracts. A. nidulans strain R153 was germinated in YG media. Total protein extracts were prepared in HK buffer in the absence of phosphatase inhibitors. Since the R153 cultures from which the protein lysates were made was asynchronous, <5% of the cells would be in mitosis. Since NIMA levels are very low throughout much of interphase, a total of 6 mg of total protein sample was used for each of the following co-immunoprecipitation experiments.

NIMA was immunoprecipitated using a NIMA specific antibody.

Immunoprecipitates were washed and then subjected to SDS-PAGE followed by western blotting with α-TINC or NIMA antibodies. NIMA was specifically immunoprecipitated using a NIMA specific antibody (Figure 6.1A NIMA Ab +

TINC Pep., NIMA Ab), but not with Pre-immune sera (Figure 6.1A Pre-Immune

Sera) or streptavidin coated paramagnetic beads alone (Figure 6.1A Beads alone). Western blotting of these immunoprecipitates with α-TINC antibody revealed that TINC was present in NIMA immunoprecipites (Figure 6.1A NIMA

Ab), and that this binding was competed by the addition of a TINC specific peptide. TINC was not present in precipitates when NIMA was absent (Figure

6.1A Pre-Immune sera, Beads alone).

162 Initial attempts to co-immunoprecipitate NIMA and TINC proved troublesome. Initially, protein lysates were made in complete HK buffer which includes an array of phosphatase inhibitors, including microcystin, which are required to maintain maximal NIMA kinase activity (Ye et al., 1995). The Two- hybrid interaction data for TINC and NIMA suggested that the interaction may be stabilized in the presence of inactive NIMA. Therefore, to examine the effect of

NIMA activity on the interaction total protein lysates were prepared from R153 in either HK buffer with phosphatase inhibitors or HK buffer with no phosphatase inhibitors. NIMA immunoprecipitates were performed as described above. TINC was detected only in NIMA immunoprecipitates prepared in the absence of phosphatase inhibitors (Figure 6.1B Compare Lane 1 to Lane 3). The addition of microcystin to lysates is known to be centrally important to maintain NIMA kinase activity. To examine the affect of microcystin on the TINC-NIMA interaction,

R153 lysates were prepared either in HK buffer with all phosphatase inhibitors

(including microsystin) or HK buffer with phosphatase inhibitors except microcystin. TINC was detected in NIMA immunoprecipitates prepared in the absence but not the presence of microcytin (Figure 6.1C). To further examine the affect of NIMA kinase activity on the interaction, protein lysates were produced from a strain containing the temperature sensitive nimA7 allele

(SO117). nimA7 is a temperature sensitive allele of nimA which displays markedly lower kinase activity at non-permissive temperature (Pu et al., 1995).

Followng a period of germination at permissive temperature, the strain was shifted to non-permissive temperature for a period of three hours to inactivate

163 NIMA7 resulting in a G2 arrest. Total protein lysates were produced from these arrested samples in the presence or absence of microcystin, and NIMA immunoprecipitaion performed. In these samples TINC was detected in NIMA immunoprecipitates made in either the absence or presence of microcystin, although slightly less TINC was present when microcystin was added (Figure

6.1D). The combination of these studies argues that the NIMA-TINC interaction is stabilized in the presence of the NIMA kinase when it is not fully activated.

In the absence of a functional anti-TIND antibody, attempts were made to co-immunoprecipitate NIMA and TIND from strains which expressed either TIND-

HA or ∆N-TIND-HA. Due to the fact that TIND interacted selectively with kinase active NIMA in the Two-hybrid screen, total protein lysates were prepared from these strains in both the absence and presence of phosphatase inhibitors.

However, NIMA and TIND failed to co-immunoprecipitate.

6.2.2. Expression of ∆N-TINC produces NIMA instability

Due to the interaction between NIMA and TINC and the cell cycle specific defects produced by ∆N-TINC and ∆N-TIND expression we examined NIMA in mitotic samples with and without ∆N-TINC or ∆N-TIND expression. These experiments make use of a temperature sensitive allele of the A. nidulans homologue of the cdc25 phosphatase, nimT23. Incubation of cells containing nimT23 at non-permissive temperature results in a G2 arrest (identified as G2 in

Figure 6.2) with inactive CDK1. Downshift to permissive temperature results in a release of cells into mitosis. Mitotic samples can be produced by the addition of

164 the microtubule destabilizer nocodoazole, to G2 arrested samples with subsequent release into permissive temperature. The addition of nocodazole activates the Spindle Mitotic Checkpoint which arrests cells in metaphase. For the following study, time course samples following release into nocodazole were produced at twenty, forty, and sixty minutes following release of cells into mitosis

(identified as 20, 40, and 60 in Figure 6.2)

Interphase and mitotic time course samples were generated as described above for vector control cells (D23C), ∆N-TINC expressing cells (JD161), and

∆N-TIND expressing cells (JD162). NIMA was immunoprecipitated from these samples using an anti-NIMA antibody, subjected to SDS-PAGE, and western blotting performed with NIMA specific antibodies. Also for each of these samples

200 µg of protein lysate was subjected to SDS-PAGE and western blotting with

NIMECyclinB antibodies to confirm cell cycle stage, and anti-tubulin antibodies

(B512) as a control for sample loading.

As expected vector control G2 samples contained a single form of NIMA

(Figure 6.2 Control G2) phosphorylated forms of NIMA with retarded mobility were obvious at 40 and 60 minutes post release into nocodazole (Figure 6.2

Control 40 and 60) as NIMA is activated and cells enter mitosis. Samples from cells expressing ∆N-TIND appeared similar to control cells (Figure 6.2 ∆N-TIND).

While ∆N-TINC expressing cells contained a single form of NIMA in G2 samples similar to control cells (Figure 6.2 ∆N-TINC G2). Interestingly, NIMA disappears from the samples which were released into nocodazole (Figure 6.2 ∆N-TINC 20-

60), suggesting that NIMA is prematurely destabilized during mitosis in these

165 cells. Strikingly, NIME levels fall sharply at 60 minutes following release suggesting that these cells exit mitosis following loss of NIMA (Figure 6.2 ∆N-

TINC 60).

6.3. Discussion

6.3.1. TINC-NIMA Interaction

The ability to co-immunoprecipitate endogenous TINC and NIMA in A. nidulans confirms the interaction observed between NIMA and TINC in the Two- hybrid screen. Additionally, these experiments suggest that the interaction between NIMA and TINC is phosphorylation state specific, which was also indicated by the selective interaction of TINC with only kinase negative forms of

NIMA in the yeast Two-hybrid screen. Initial TINC-NIMA interaction experiments demonstrated that addition of the phosphatase inhibitor microcystin to protein extracts destabilized the interaction. NIMA is known to be far less active in the absence of microcystin (Ye et al., 1995). This fact suggests that the NIMA-TINC interaction is stabilized in the presence of inactive kinase. Additionally, the fact that TINC was able to be co-immunoprecipitated with a kinase inactivated allele of NIMA (nimA7) at restrictive temperature in both the absence and presence of phosphatase inhibitors, further supports the idea that the interaction is stabilized in the presence of inactive NIMA. The nature of the TINC-NIMA interaction suggests that TINC and NIMA may interact most strongly during G2 at a time when NIMA is not fully activated. Alternately, TINC may represent a NIMA

166 substrate. In this case the transient enzyme substrate interaction between NIMA and TINC may be stabilized in the presence of an inactive kinase allowing for the detection of this interaction in both genetic and biochemical studies.

6.3.2. Premature loss of NIMA

The ability of ∆N-TINC to promote loss of NIMA from mitotic extracts is intriguing in that it suggests a possible mechanism through which ∆N-TINC acts to promote cell cycle defects. As discussed previously, NIMA must exist at a level sufficient for mitotic initiation during late interphase in cells expressing ∆N-

TINC. The premature loss of NIMA, although not examined directly previously, would be expected to produce mitotic defects. The generation of spindle and nuclear envelope defects would agree well with spindle specific roles suggested by the interaction between NIMA and the SPB associated protein, TINA (Osmani et al., 2003), the localization of NIMA to components of the mitotic spindle during mitosis (De Souza et al., 2000), and the spindle defects observed with overexpression of NIMA (Osmani et al., 1988b). The inability of ∆N-TIND to induce loss of NIMA may be due to insufficient levels of ∆N-TIND expression.

6.3.3. A TIND-NIMA interaction in A. nidulans?

The inability to show that NIMA and TIND co-immunoprecipitate in A. nidulans while disappointing was not unexpected. The TIND localization studies

I have presented convincingly reveal TIND to be a mitochondrial protein. Past

NIMA localization studies have suggested no role for NIMA at the mitochondria.

167 These studies do not, however, rule out a potential interaction between NIMA and TIND. As suggested previously, a small amount of TIND, undetectable through the immunofluorescence procedures outlined in this study, may be present outside of the mitochondria and available for interaction with NIMA. The additional possibility that non-mitochondrial forms of TIND may exist under conditions of cellular stress was also addressed in Chapter 3.

The inability to demonstrate an interaction between NIMA and ∆N-TIND was more surprising, since this form is present outside of the mitochondria and produces cell cycle specific defects. The inability to co-immunoprecipitate NIMA and ∆N-TIND in these studies does not rule out the possibility that they do in fact interact. The level of ∆N-TIND expression may not be high enough in these cells to detect the interaction. Alternately, ∆N-TIND and NIMA may not interact and

∆N-TIND may induce the defects seen in a manner independent from that used by ∆N-TINC (inhibition of NIMA), perhaps through direct perturbation of spindle function.

168

Figure 6.1. TINC binds NIMA in A. nidulans. (A) Total protein lysates were prepared in the absence of phosphatase inhibitors from strain R153 grown at

32°C to log phase. NIMA was immunoprecipitated with anti-NIMA antibodies.

NIMA immunoprecipitates were subjected to SDS-PAGE and western blotting with α-TINC antibodies. (B) The NIMA-TINC interaction is phosphorylation state sensitive. Protein was prepared from strain R153 in the absence (Phosphatase

Inh. -) or presence (Phosphatase Inh. +) of phosphatase inhibitors. NIMA immunoprecipitates were probed for TINC. (C and D) Total protein lysates were prepared in the presence of phosphatase inhibitors either with or without microcystin from (C) R153 or (D) a strain containing the nimA7 temperature sensitive allele (SO117). Both strains were grown at 30°C to mid log phase and then shifted to 42°C to inactivate NIMA (in SO117). After three hours at 42°C strainwere harvested and protein lysates prepared. NIMA immunoprecipitates were probed for TINC.

169 . a p r A e C e s P a e C r e n n N e I u lo s b T t a e e u A m + n n p s im u lo n A - d b b I a m a M e A A g I r e s im N P B A A - d m M M e a 0 I I r e 0 N N P B 2 NIMA

NIMA

TINC

+ microcystin - microcystin

TINC

a D r e s e e Lanes 1 2 3 4 n n B u lo b m a A NIMA Ab m s - - A i + + - d M e a Phosphatase Inh I r e - - + + N P B

NIMA NIMA

TINC TINC + microcystin - microcystin

170 Control ∆N-TIND ∆N-TINC

G2 20 40 60 G2 20 40 60 G2 20 40 60

NIMA

NIMECyclinB

Tubulin

Figure 6.2. Cells expressing ∆N-TINC prematurely lose NIMA. Cells expressing ∆N-TINC-HA (JD161), ∆N-TIND-HA (JD162), or control cells (D23C) were germinated in minimal media yeast lactose with 40 mM threonine at 30°C to early log phase. Cells were then rapidly shifted to 42°C for three hours to inactivate the temperature nimt23 allele and accumulate cells in G2 (G2). Cells were subsequently release into mitosis by downshifting cultures to 30°C in the presence of nocodazole. Time course samples were removed at 20 (20), 40 (40), and 60 (60) minutes following release from non-restrictive temperature. NIMA was immunoprecipitated using anti-NIMA antibodies from 5 mg of protein sample at each time point and subjected to SDS-PAGE and western blotting with anti-

NIMA antibodies (NIMA). An additional 200µg of each sample was also subjected to SDS-PAGE and western blotting with anti-NIMECyclinB (NIME) and anti-tubulin antibodies (Tubulin).

171

CHAPTER 7

FINAL DISCUSSION AND FUTURE DIRECTIONS

7.1 Overview

The aim of this project was to identify and characterize NIMA interacting proteins in A. nidulans. Activation of the NIMA protein kinase is absolutely required for all aspects of mitosis in A. nidulans. We have used a yeast Two- hybrid screen utilizing different forms of NIMA as bait to screen a library of A. nidulans cDNAs. Six cDNAs encoding potential NIMA interacting proteins were identified in this screen, tinA through tinF. Overexpression of either tinC or tinD produced a cell cycle specific defect in A. nidulans which prompted the further study of these two proteins.

TINC is a member of a fungal specific family of proteins and shares a high degree of homology with N. crassa Het-C. Sequencing of the tinC locus in twenty-five strains of A. nidulans identified no multi-allelism. Additionally, a

TINC-like protein, An-HETC was identified in A. nidulans which shares a higher degree of homology with Het-C, however, An-hetC also appears to lack multi- allelism. Together these data suggested that TINC is not involved in heterokaryon incompatibility in A. nidulans. TINC is a cytoplasmic protein which

172 is also present in the nucleus during mitosis. Deletion analysis of tinC alone or in combination with An-hetC demonstrated that tinC and An-hetC functions are non- essential under the conditions tested, or that an additional layer of functional redundancy exists in A. nidulans. Sequencing of the tinC cDNA recovered from the Two-hybrid screen revealed it was 5’ truncated so a full-length clone was obtained and expressed in A. nidulans to determine whether they produced cell cycle defects. Disappointingly, expression of full-length TINC failed to produce cell cycle specific defects. This difference may be due in part to the altered localization of ∆N-TINC compared to the full-length protein. ∆N-TINC accumulates at the periphery of membranes, which at high expression levels associate with nuclei. Cells which express ∆N-TINC proceed through faulty mitoses. These cells also displayed striking nuclear membrane fission defects as even cells with separated masses of DNA frequently contained a single highly stretched nucleus. TINC and NIMA interact in A. nidulans in a phosphorylation specific manner, with TINC apparently interacting with inactive forms of NIMA.

Finally, expression of ∆N-TINC produced a premature loss of NIMA during a mitotic arrest, suggesting a possible mechanism for ∆N-TINC defects.

TIND is a highly conserved protein with homologues in bacteria through to humans. All TIND homologues contain a carboxy-terminal domain which is highly identical to the carboxy-terminal domain of NIFU, the function of which is unknown. TIND is a mitochondrial protein throughout the cell cycle. ∆N-TIND is present throughout the cell. Expression of both TIND and ∆N-TIND inhibits colony formation but only ∆N-TIND expression produced cell cycle specific

173 defects. Similar to ∆N-TINC, cells which express ∆N-TIND undergo faulty mitoses and display nuclear membrane fission defects. Unlike ∆N-TINC, ∆N-

TIND expression was typified by the presence of aberrant mitotic spindles which resembled monopolar spindles.

7.2. A potential role for TINC in cell cycle regulation

The interaction between TINC and NIMA both in the Two-hybrid screen and in A. nidulans coupled with the ability of ∆N-TINC expression to produce striking cell cycle specific defects, strongly suggests a role for TINC in cell cycle regulation in A. nidulans. The failure of the full-length TINC to produce similar defects when expressed in A. nidulans was disappointing and leaves open the question whether defects produced by ∆N-TINC expression are a reflection of a dominant negative or gain of function phenotype. Basically, are these defects a reflection of the role of endogenous TINC or do they represent defects in a pathway in which TINC does not normally participate? It was hoped that deletion of tinC or tinC in combination with An-hetC would result in cell cycle progression or nuclear envelope defects, however the lack of either of these phenotypes does not allow us to say with certainty that ∆N-TINC phenotypes represent the expression of a dominant negative form of TINC, it does not however rule out this possibility. In fact the evolutionary conservation of two tinC / An-hetC homologues in filamentous fungi argues for TINC fulfilling an important, potentially cell cycle specific, function in A. nidulans.

174 The interaction between TINC and NIMA in A. nidulans represents a strong indication that TINC does possess a cell cycle specific role. The premature loss of NIMA in mitotically arrested samples expressing ∆N-TINC suggests a possible role for TINC in NIMA stabilization in mitosis. This is an attractive role as, NIMA levels are known to be tightly regulated at the levels of mRNA abundance, protein levels, phosphorylation and proteolytic degradation.

The central role for NIMA during mitosis suggests that some of these regulatory functions would almost certainly be redundant, providing a possible explanation for the lack of any striking ∆tinC phenotypes. This would also provide an explanation for the lack of any phenotypes observed for TINC overexpression.

For example the stabilization of NIMA during interphase may not produce any striking defects since NIMA activity is also strictly regulated at the level of phosphorylation and subcellular localization.

To determine the role of TINC in A. nidulans, will require an expansion of the limited search for synthetic lethality between ∆tinC/∆hetC and the limited number of cell cycle regulatory mutants described here. The identification and characterization of these genetic interactors should provide insight into the role of

TINC and may illuminate additional levels of functional redundancy. The level of evolutionary conservation of TINC among filamentous fungi, coupled with the interaction between NIMA and TINC and the striking defects produced by expression of truncated forms of TINC warrant further study of TINC.

175 7.3. A search for new alleles of nimA

The original screen by Ron Morris identified four temperature sensitive nimA alleles. All of these mutations produce an arrest in G2, emphasizing the essential role for NIMA at mitotic initiation, however, several pieces of information suggest a continuing role for NIMA during mitosis. First, NIMA activity peaks at the G2-M transition, but persists throughout mitosis. NIMA localizes to the spindle apparatus during mitosis and interacts with a spindle pole body associated protein, TINA (De Souza et al. 2000; Osmani et al. 2003). Cells which enter mitosis with inadequate NIMA in the presence of an inactivated component of the APC, BIME7, (Osmani et al., 1991b) exhibit severe defects in nuclear envelope and spindle morphologies. Finally, this study supports a mitotic specific role for NIMA as premature loss of NIMA correlates with nuclear membrane fission defects. The results of these earlier studies coupled with those described here represent a strong suggestion that the identification of additional alleles of nimA may be worthwhile. For example mutations resulting in a frame shift mutation or premature STOP which would eliminate the PEST sequence from the carboxy-terminus of NIMA would produce bim phenotype as cells which express non-degradable NIMA have been shown to arrest in mitosis

(Pu and Osmani, 1995). Additional nimA alleles may allow mitotic entry but produce mitotic defects including spindle defects and nuclear membrane fission defects. These mutations may either produce a mitotic arrest or allow mitotic exit, as suggested by the studies described here. Since many of these phenotypes would be lethal, conditional mutations would need to be generated.

176 7.4. Two models addressing ∆N-TINC and ∆N-TIND defects

One of the central themes of this thesis is the ability of expression of two vastly different proteins to induce strikingly similar cell cycle defects in A. nidulans. I will discuss two potential explanations for this phenomenon. First, the defects are a reflection of NIMA function with TINC and TIND acting as NIMA inhibitors. Second, expression of TINC and TIND activate a common novel checkpoint mechanism.

First, the common denominator between TINC and TIND is that they were both isolated as NIMA interacting proteins. The inability to confirm the NIMA-

TIND interaction in A. nidulans while disappointing, represents negative data which does not preclude the possibility that TIND and or ∆N-TIND interact with

NIMA. The striking similarity of the defects induced by ∆N-TINC or ∆N-TIND also supports a common target (NIMA). If we assume that ∆N-TINC and ∆N-

TIIND function essentially as NIMA inhibitory proteins which result in premature destabilization of NIMA (as suggested by ∆N-TINC overexpression), it is possible to model the defects in nuclear membrane fission we observed based on this inhibition of NIMA function given new information which has emerged from our laboratory.

Recent work in our lab has suggested that one of the primary mechanisms for initiating mitosis in A. nidulans is the dramatic restructuring of the nuclear pore complex (De Souza et al., 2004 Submitted). The NIMA kinase was previously shown to interact with two nucleoporins. Examination of the localization of these two nucleoporins and others has shown that they are

177 present at the NPC during interphase, but are dispersed at mitosis (De Souza et al., 2004 Submitted). Further studies have implicated the NIMA kinase as central to the loss of several nucleoporins from the nuclear pore at mitosis (De Souza et al., Submitted). Expression of NIMA in hydroxyurea arrested cells can induce the release of GFP-SONBNUP98 from the nuclear periphery (De Souza et al., 2004

Submitted). Notably SONB contains a large number of potential NIMA phosphorylation sites and mitotic forms with highly decreased mobility have been detected by SDS-PAGE (De Souza et al., 2003). These data suggest that a primary role of NIMA in mitotic initiation is the partial disassembly of the nuclear pore complex by inducing the release of some nucleoporins by direct phosphorylation (Figure 7.1 WT). In the first hypothesis, proteolytic destruction of NIMA at mitotic exit would allow nucleoporins to reassociate resulting in a resumption of regulated transport which characterizes interphase (Figure 7.1

WT).

In contrast, cells which express ∆N-TINC or ∆N-TIND would lose nucleoporins from the nuclear periphery and enter mitosis (Figure 7.1). During mitosis NIMA would be lost prematurely either through the direct or indirect action of ∆N-TINC or ∆N-TIND (Figure 7.1). Loss of NIMA would result in early reassociation of nucleoporins and a resumption of regulated transport producing a premature return to a G1-like nucleus (Figure 7.1). This premature return to interphase would result in the failure to complete normal mitotic exit functions including nuclear membrane fission (Figure 7.1).

178 Studies performed recently in our lab support the idea that a loss of NIMA would allow nucleoporin reassociation. Expression of NIMA in S-phase arrested cells results in a release of SONB from the nuclear periphery. This release is followed by a cycling of reassociation and disassociation of SONB (De Souza et al. 2004 Submitted). This oscillatory behavior was reminiscent of fluctuations in

NIMA following a rapid inactivation of the APC component BIMA (Ye et al., 1998).

In support of this idea, expression of stable forms of NIMA results in a release of

SONB from the nuclear pore with no reassociation (De Souza et al., 2004

Submitted).

A second possibility is that ∆N-TINC and ∆N-TIND may generate mitotic defects through independent mechanisms but trigger a common checkpoint system which allows mitotic exit but blocks nuclear membrane fission. In support of this idea, a number of other studies have identified similar nuclear divison defects similar to those seen for ∆N-TINC and ∆N-TIND. Studies which examined the loss of NIMU, an A. nidulans Pot1 homologue, demonstrated that these cells undergo early mitotic exit. The nuclei in these cells are highly stretched, some with discernable DNA linkage between larger DNA foci (Pitt et al., 2004). Activation of DNA checkpoint machinery in cells lacking the ability to tyrosine phosphorylate CDK1 also results in the production of highly stretched polyploidy nuclei. Although these defects are generated by rereplication of DNA is the absence of intervening mitoses (De Souza et al., 1999). Finally, cells containing certain γ-tubulin mutations produce abnormal spindles and proceed

179 through faulty mitoses without undergoing nuclear division (Prigozhina et al.,

2004).

Additionally, the ability of A. nidulans to haploidize is well established.

Diploids can be induced to undergo chromosome loss in a laboratory setting.

Significantly, the cells resulting from this process appear to be perfectly viable haploids.

Basically, in the presence of mitotic defects, A. nidulans would forego nuclear membrane fission and maintain all DNA within a single nucleus. This would effectively maintain a diploid nucleus rather than undergoing nuclear membrane fission which would produce severe aneuploidy. A. nidulans would then be able to regenerate haploid nuclei through chromosome loss, which appears to be a coordinated process in A. nidulans.

7.5. ∆N- TIND and redox regulation

As mentioned previously, the carboxy-terminal domain of TIND is homologous to the carboxy-terminal domain of NIFU. This domain has been termed the thioredoxin module due to the conservation of two cysteine resdues present in the active site of thioredoxin in a similar context in this domain. In thioredoxin these cysteines participate directly in reduction reactions carried out thioredoxin. Thioredoxin in association with glutaredoxin regulates the redox state of cells in organisms from bacteria to humans (Holmgren, 1985; Holmgren,

1989). The potential that this domain is capable of carrying out reduction

180 reactions is intriguing in that it suggests a possible mechanism for ∆N-TIND generating mitotic and specifically spindle defects.

A recent study has demonstrated that microtubule assembly is altered by cysteine oxidation of two microtubule associated proteins, tau and MAP-2

(Landino et al., 2004). Potentially, misolocalization of ∆N-TIND to the cytoplasm and nucleus may represent the mislocalization of a protein capable of altering the oxidation state of proteins. For example, ∆N-TIND could alter the oxidation state of microtubule proteins and perturb microtubule assembly.

7.6. NIMA: maintaining the mitotic state and nuclear membrane fission

Note that the above models suggest an active role for NIMA in maintaining the mitotic state by blocking nucleoporin reassociation. Studies involving expression of stable forms of NIMA have also raised this possibility as these cells are unable to exit mitosis even after cyclin B is degraded (Pu and Osmani, 1995).

This mechanism would be somewhat unique from that seen in other eukaryotic systems where the primary mechanism of preventing mitotic exit is through the stabilization of cyclin B through inactivation of the APC.

The studies described in this thesis also suggest a potential role for NIMA in nuclear membrane fission. NIMA may coordinate nuclear membrane fission and mitotic exit either passively by inhibiting mitotic exit until these events are completed or actively through interactions with additional regulators of this process.

181

Figure 7.1. Control of nuclear membrane fission and mitotic exit by NIMA.

During G2 nucleoporins regulate transport at the nuclear pore complex. During mitotic entry the NIMA kinase phosphorylates nucleoporins, resulting in their release from the nuclear pore complex. Loss of nucleoporins allows mitotic specific regulators to enter the nucleus and establish mitosis. Under normal conditions NIMA activity is maintained through mitosis until mitotic exit when

NIMA is degraded. Loss of NIMA allows for reassociation of nucleoporins and reestablishment of the regulated transport which characterizes interphase.

Alternately, premature loss or inactivation of NIMA would potentially allow reassociation of nucleoporins resulting in a premature mitotic exit and failure to complete late mitotic events like nuclear membrane fission.

182 Premature NIMA Wild Type Inactivation

G2 G2 NIMA

M M

Premature NIMA degradation & mitotic exit NIMA M M G1 G1

NIMA degradation & mitotic exit NIMA G1 G1 G1 G1

= phosphorylation

= nuclear envelope = nucleoporins

= karyofission

183

BIBLIOGRAPHY

Agar,J.N., Yuvaniyama,P., Jack,R.F., Cash,V.L., Smith,A.D., Dean,D.R., and Johnson,M.K. (2000). Modular organization and identification of a mononuclear iron- within the NifU protein. J. Biol. Inorg. Chem. 5, 167-177.

Aleksenko,A. and Clutterbuck,A.J. (1997). Autonomous plasmid replication in Aspergillus nidulans: AMA1 and MATE elements. Fungal. Genet. Biol. 21, 373-387.

Aleksenko,A., Nikolaev,I., Vinetski,Y., and Clutterbuck,A.J. (1996). Gene expression from replicating plasmids in Aspergillus nidulans. Mol. Gen. Genet. 253, 242-246.

Anwar,M.M., Croft,J.H., and Dales,R.B. (1993). Analysis of heterokaryon incompatibility between heterokaryon-compatibility (H-c) groups R and GL provides evidence that at least eight het loci control somatic incompatibility in Aspergillus nidulans. J. Gen. Microbiol. 139, 1599-1603.

Askjaer,P., Galy,V., Hannak,E., and Mattaj,I.W. (2002). Ran GTPase cycle and importins alpha and beta are essential for spindle formation and nuclear envelope assembly in living Caenorhabditis elegans embryos. Mol. Biol. Cell 13, 4355-4370.

Bailey,C. and Arst,H.N., Jr. (1975). Carbon catabolite repression in Aspergillos nidulans. Eur. J. Biochem. 51, 573-577.

Ballance,D.J. and Turner,G. (1985). Development of a high-frequency transforming vector for Aspergillus nidulans. Gene 36, 321-331.

Beach,D., Durkacz,B., and Nurse,P. (1982). Functional homologous cell cycle control genes in budding and fission yeast. Nature 300, 705-709.

Beaudouin,J., Gerlich,D., Daigle,N., Eils,R., and Ellenberg,J. (2002). Nuclear envelope breakdown proceeds by microtubule-induced tearing of the lamina. Cell 108, 83-96.

Bergen,L.G. and Morris,N.R. (1983). Kinetics of the nuclear division cycle of Aspergillus nidulans. J. Bacteriol. 156, 155-160. 184 Bergen,L.G., Upshall,A., and Morris,N.R. (1984). S-phase, G2, and nuclear division mutants of Aspergillus nidulans. J. Bacteriol. 159, 114-119.

Bharadwaj,R. and Yu,H. (2004). The spindle checkpoint, aneuploidy, and cancer. Oncogene 23, 2016-2027.

Bird,D. and Bradshaw,R. (1997). Gene targeting is locus dependent in the filamentous fungus Aspergillus nidulans. Mol. Gen. Genet. 255, 219-225.

Blake,D.J., Tinsley,J.M., Davies,K.E., Knight,A.E., Winder,S.J., and Kendrick- Jones,J. (1995). Coiled-coil regions in the carboxy-terminal domains of dystrophin and related proteins: potentials for protein-protein interactions. Trends Biochem. Sci. 20, 133-135.

Boehmer,T., Enninga,J., Dales,S., Blobel,G., and Zhong,H. (2003). Depletion of a single nucleoporin, Nup107, prevents the assembly of a subset of nucleoporins into the nuclear pore complex. Proc Natl Acad Sci U. S. A 100, 981-985.

Booher,R. and Beach,D. (1987). Interaction between cdc13+ and cdc2+ in the control of mitosis in fission yeast; dissociation of the G1 and G2 roles of the cdc2+ protein kinase. EMBO J. 6, 3441-3447.

Booher,R. and Beach,D. (1988). Involvement of cdc13+ in mitotic control in Schizosaccharomyces pombe: possible interaction of the gene product with microtubules. EMBO J. 7, 2321-2327.

Burgess,B.K. and Lowe,D.J. (1996). Mechanism of molybdenum nitrogenase. Chem. Rev. 96, 2983-3012.

Caten,C.E. (1972). Vegetative incompatibility and cytoplasmic infection in fungi. J. Gen. Microbiol. 72, 221-229.

Champe,S.P. and Simon,L.D. (1992). Cellular differentiation and tissue formation in fungus Aspergillus nidulans. In Morphogenesis: An analysis of the development of biological structure, E.F.Rossomando and S.Alexander, eds. (New York: Marcel Dekker, Inc.), pp. 63-91.

Chaveroche,M.K., Ghigo,J.M., and d'Enfert,C. (2000). A rapid method for efficient gene replacement in the filamentous fungus Aspergillus nidulans. Nuc. Acids Res. 28, E97.

Chen,Y., Riley,D.J., Zheng,L., Chen,P.L., and Lee,W.H. (2002). Phosphorylation of the mitotic regulator protein hec1 by nek2 kinase is essential for faithful chromosome segregation. J. Biol. Chem. 277, 49408-49416.

185 Cheng,J., Park,T.S., Chio,L.C., Fischl,A.S., and Ye,X.S. (2003). Induction of apoptosis by sphingoid long-chain bases in Aspergillus nidulans. Mol. Cell Biol. 23, 163-177.

Chiu,A., Revenkova,E., and Jessberger,R. (2004). DNA interaction and dimerization of eukaryotic SMC hinge domains. J. Biol. Chem.

Ciosk,R., Shirayama,M., Shevchenko,A., Tanaka,T., Toth,A., Shevchenko,A., and Nasmyth,K. (2000). Cohesin's binding to chromosomes depends on a separate complex consisting of Scc2 and Scc4 proteins. Mol. Cell 5, 243- 254.

Ciosk,R., Zachariae,W., Michaelis,C., Shevchenko,A., Mann,M., and Nasmyth,K. (1998). An ESP1/PDS1 complex regulates loss of sister chromatid cohesion at the metaphase to anaphase transition in yeast. Cell 93, 1067- 1076.

Claros,M.G. and Vincens,P. (1996). Computational method to predict mitochondrially imported proteins and their targeting sequences. Eur. J. Biochem. 241, 779-786.

Creaser,E.H., Porter,R.L., Britt,K.A., Pateman,J.A., and Doy,C.H. (1985). Purification and preliminary characterization of alcohol dehydrogenase from Aspergillus nidulans. Biochem. J. 225, 449-454.

Cuddihy,A.R. and O'Connell,M.J. (2003). Cell-cycle responses to DNA damage in G2. Int. Rev. Cytol. 222, 99-140.

Dales,R.B., Moorhouse,J., and Croft,J.H. (1993). Evidence for a multi-allelic heterokaryon incompatibility (het) locus detected by hybridization among three heterokaryon-compatibility (h-c) groups of Aspergillus nidulans. Heredity 70, 537-543.

Dales,R.B. and Croft,J.H. (1990). Investigation of the het genes that control heterokaryon incompatibility between members of heterokaryon- compatibility (h-c) groups A and G1 of Aspergillus nidulans. J. Gen. Microbiol. 136, 1717-1724.

De Souza,C.P., Horn,K.P., Masker,K., and Osmani,S.A. (2003). The SONB(NUP98) Nucleoporin Interacts With the NIMA Kinase in Aspergillus nidulans. Genetics 165, 1071-1081.

De Souza,C.P., Osmani,A.H., Wu,L.P., Spotts,J.L., and Osmani,S.A. (2000). Mitotic histone H3 phosphorylation by the NIMA kinase in Aspergillus nidulans. Cell 102, 293-302.

186 De Souza,C.P.C., Ye,X.S., and Osmani,S.A. (1999). Checkpoint defects leading to premature mitosis also cause endoreplication of DNA in Aspergillus nidulans. Mol. Biol. Cell 10, 3661-3674.

DeLuca,J.G., Howell,B.J., Canman,J.C., Hickey,J.M., Fang,G., and Salmon,E.D. (2003). Nuf2 and Hec1 are required for retention of the checkpoint proteins Mad1 and Mad2 to kinetochores. Curr. Biol. 13, 2103-2109.

Doonan,J.H. (1992). Cell division in Aspergillus. J. Cell Sci. 103, 599-611.

Doonan,J.H., MacKintosh,C., Osmani,S., Cohen,P., Bai,G., Lee,E.Y.C., and Morris,N.R. (1991). A cDNA encoding rabbit muscle protein phosphatase 1a complements the Aspergillus cell cycle mutation, bimG11. J. Biol. Chem. 266, 18889-18894.

Dou,X., Wu,D., An,W., Davies,J., Hashmi,S.B., Ukil,L., and Osmani,S.A. (2003). The PHOA and PHOB Cyclin-Dependent Kinases Perform an Essential Function in Aspergillus nidulans. Genetics 165, 1105-1115.

Draetta,G., Luca,F., Westendorf,J., Brizuela,L., Ruderman,J., and Beach,D. (1989). Cdc2 protein kinase is complexed with both cyclin A and B: evidence for proteolytic inactivation of MPF. Cell 56, 829-838.

Eidam,E. (1883). Zur Kenntniss der Entwicklung bei den Ascomyceten. III Sterigmatocystis nidulans. FS Cohn, Beitr Biol Pflanzen 3, 392-441.

Evans,T.E., Rosenthal,J., Youngbloom,K., Distel,K., and Hunt,T. (1983). Cyclin: a protein specified by maternal mRNA in sea urchin eggs that is destroyed at each cleavage division. Cell 33, 389-396.

Faragher,A.J. and Fry,A.M. (2003). Nek2A kinase stimulates centrosome disjunction and is required for formation of bipolar mitotic spindles. Mol. Biol. Cell 14, 2876-2889.

Fiddy,C. and Trinci,A.P. (1976). Mitosis, septation, branching and the duplication cycle in Aspergillus nidulans. J. Gen. Microbiol. 97, 169-184.

Fields,S. and Song,O. (1989). A novel genetic system to detect protein-protein interactions. Nature 340, 245-246.

Frazzon,J., Fick,J.R., and Dean,D.R. (2002). Biosynthesis of iron-sulphur clusters is a complex and highly conserved process. Biochem. Soc. Trans. 30, 680-685.

187 Fry,A.M., Mayor,T., Meraldi,P., Stierhof,Y.D., Tanaka,K., and Nigg,E.A. (1998a). C-Nap1, a novel centrosomal coiled-coil protein and candidate substrate of the cell cycle-regulated protein kinase Nek2. J. Cell. Biol. 141, 1563- 1574.

Fry,A.M., Meraldi,P., and Nigg,E.A. (1998b). A centrosomal function for the human Nek2 protein kinase, a member of the NIMA family of cell cycle regulators. EMBO J. 17, 470-481.

Ganesh,S., Tsurutani,N., Suzuki,T., Ueda,K., Agarwala,K.L., Osada,H., Delgado- Escueta,A.V., and Yamakawa,K. (2003). The Lafora disease gene product laforin interacts with HIRIP5, a phylogenetically conserved protein containing a NifU-like domain. Hum. Mol. Genet. 12, 2359-2368.

Garnjobst,L. and Wilson,J.F. (1956). Heterokaryon and protoplasmic incompatibility in Neurosporra crassa. Proc. Natl. Acad. Sci. U. S. A. 42, 613-618.

Gautier,J., Norbury,C., Lohka,M., Nurse,P., and Maller,J. (1988). Purified maturation-promoting factor contains the product of a Xenopus homolog of the fission yeast cell cycle control gene cdc2+. Cell 54 , 433-439.

Gonczy,P. (2002). Nuclear envelope: torn apart at mitosis. Curr. Biol. 12, R242- R244.

Gordon,C.L., Khalaj,V., Ram,A.F., Archer,D.B., Brookman,J.L., Trinci,A.P., Jeenes,D.J., Doonan,J.H., Wells,B., Punt,P.J., van den Hondel,C.A., and Robson,G.D. (2000). Glucoamylase::green fluorescent protein fusions to monitor protein secretion in Aspergillus niger. Microbiology 146, 415-426.

Gould,K.L. and Nurse,P. (1989). Tyrosine phosphorylation of the fission yeast cdc2+ protein kinase regulates entry into mitosis. Nature 342, 39-45.

Grallert,A. and Hagan,I.M. (2002). Schizosaccharomyces pombe NIMA-related kinase, Fin1, regulates spindle formation and an affinity of Polo for the SPB. EMBO J. 21, 3096-3107.

Grallert,A., Krapp,A., Bagley,S., Simanis,V., and Hagan,I.M. (2004). Recruitment of NIMA kinase shows that maturation of the S. pombe spindle-pole body occurs over consecutive cell cycles and reveals a role for NIMA in modulating SIN activity. Genes Dev. 18, 1007-1021.

Grindle,M. (1963a). Heterokaryon compatibility of closely related wild isolates of Aspergillus nidulans. Heredity 18, 397-405.

Grindle,M. (1963b). Heterokaryon compatibility of unrelated strains in the Aspergillus nidulans group. Heredity 18, 191-204. 188 Guo,X.W., Th'ng,J.P.H., Swank,R.A., Anderson,H.J., Tudan,C., Bradbury,E.M., and Roberge,M. (1995). Chromosome condensation induced by fostriecin does not require p34cdc2 kinase acitivity and histone H1 hyperphosphorylation, but is associated with enchanced histone H2A and H3 phosphorylation. EMBO J. 14, 976-985.

Hachet,V., Kocher,T., Wilm,M., and Mattaj,I.W. (2004). Importin alpha associates with membranes and participates in nuclear envelope assembly in vitro. EMBO J. 23, 1526-1535.

Haering,C.H., Lowe,J., Hochwagen,A., and Nasmyth,K. (2002). Molecular architecture of SMC proteins and the yeast cohesin complex. Mol. Cell 9, 773-788.

Harel,A., Chan,R.C., Lachish-Zalait,A., Zimmerman,E., Elbaum,M., and Forbes,D.J. (2003a). Importin beta negatively regulates nuclear membrane fusion and nuclear pore complex assembly. Mol. Biol. Cell 14, 4387-4396.

Harel,A., Orjalo,A.V., Vincent,T., Lachish-Zalait,A., Vasu,S., Shah,S., Zimmerman,E., Elbaum,M., and Forbes,D.J. (2003b). Removal of a single pore subcomplex results in vertebrate nuclei devoid of nuclear pores. Mol. Cell 11, 853-864.

Harris,S.D., Hamer,L., Sharpless,K.E., and Hamer,J.E. (1997). The Aspergillus nidulans sepA gene encodes an FH1/2 protein involved in cytokinesis and the maintenance of cellular polarity. EMBO J. 16, 3474-3483.

Harris,S.D., Morrell,J.L., and Hamer,J.E. (1994). Identification and characterization of Aspergillus nidulans mutants defective in cytokinesis. Genetics 136, 517-532.

Hartwell,L.H., Culotti,J., Pringle,J.R., and Reid,B.J. (1974). Genetic control of the cell division cycle in yeast. Science 183, 46-51.

Hartwell,L.H. and Weinert,T.A. (1989). Checkpoints: controls that ensure the order of cell cycle events. Science 246, 629-634.

Hashimoto,Y., Akita,H., Hibino,M., Kohri,K., and Nakanishi,M. (2002). Identification and characterization of Nek6 protein kinase, a potential human homolog of NIMA histone H3 kinase. Biochem. Biophys. Res. Commun. 293, 753-758.

Heichman,K.A. and Roberts,J.M. (1994). Rules to replicate by. Cell 79, 557-562.

189 Hendzel,M.J., Wei,Y., Mancini,M.A., Van Hooser,A., Ranalli,T., Brinkley,B.R., Bazett-Jones,D.P., and Allis,C.D. (1997). Mitosis-specific phosphorylation of histone H3 initiates primarily within pericentromeric heterochromatin during G2 and spreads in an ordered fashion coincident with mitotic chromosome condensation. Chromosoma 106, 348-360.

Holland,P.M., Milne,A., Garka,K., Johnson,R.S., Willis,C.R., Sims,J.E., Rauch,C.T., Bird,T.A., and Virca,G.D. (2002). Purification, cloning and characterization of Nek8, a novel NIMA- related kinase, and its candidate substrate Bicd2. J. Biol. Chem. 277, 16229-16240.

Holmgren,A. (1985). Thioredoxin. Annu. Rev. Biochem. 54, 237-271.

Holmgren,A. (1989). Thioredoxin and glutaredoxin systems. J. Biol. Chem. 264, 13963-13966.

Hori,T., Haraguchi,T., Hiraoka,Y., Kimura,H., and Fukagawa,T. (2003). Dynamic behavior of Nuf2-Hec1 complex that localizes to the centrosome and centromere and is essential for mitotic progression in vertebrate cells. J. Cell Sci. 116, 3347-3362.

Howlett,B.J., Leslie,J.F., and Perkins,D.D. (1993). Putative multiple alleles at the vegetative (heterokaryon) incompatibility loci het-c and het-8 in Neurospora crassa. Fungal Genetics Newsletter 40, 40-42.

Jacobson,D.J., Beurkens,K., and Klomparens,K.L. (1998). Microscopic and Ultrastructural Examination of Vegetative Incompatibility in Partial Diploids Heterozygous at het Loci in Neurospora crassa. Fungal Genet. Biol. 23, 45-56.

Jinks,J.L. and Grindle,M. (1963). The genetical basis of heterokaryon incompatibility in Aspergillus nidulans. Heredity 18, 407-411.

Kandli,M., Feige,E., Chen,A., Kilfin,G., and Motro,B. (2000). Isolation and characterization of two evolutionarily conserved murine kinases (Nek6 and nek7) related to the fungal mitotic regulator, NIMA. Genomics 68, 187-196.

Ke,Y.W., Dou,Z., Zhang,J., and Yao,X.B. (2003). Function and regulation of Aurora/Ipl1p kinase family in cell division. Cell Res. 13, 69-81.

Khalaj,V., Brookman,J.L., and Robson,G.D. (2001). A study of the protein secretory pathway of Aspergillus niger using a glucoamylase-GFP fusion protein. Fungal Genet. Biol. 32, 55-65.

Krien,M.J., Bugg,S.J., Palatsides,M., Asouline,G., Morimyo,M., and O'Connell,M.J. (1998). A NIMA homologue promotes chromatin condensation in fission yeast. J. Cell Sci. 111, 967-976. 190 Krien,M.J., West,R.R., John,U.P., Koniaras,K., McIntosh,J.R., and O'Connell,M.J. (2002). The fission yeast NIMA kinase Fin1p is required for spindle function and nuclear envelope integrity. EMBO J. 21, 1713-1722.

Labbe,J.C., Capony,J.P., Caput,D., Cavadore,J.C., Derancourt,J., Kaghad,M., Lelias,J.M., Picard,A., and Doree,M. (1989a). MPF from starfish oocytes at first meiotic metaphase is a heterodimer containing one molecule of cdc2 and one molecule of cyclin B. EMBO J. 8, 3053-3058.

Labbe,J.C., Picard,A., Peaucellier,G., Cavadore,J.C., Nurse,P., and Doree,M. (1989b). Purification of MPF from starfish: identification as the H1 histone kinase p34cdc2 and a possible mechanism for its periodic activation. Cell 57, 253-263.

Lamour,V., Lecluse,Y., Desmaze,C., Spector,M., Bodescot,M., Aurias,A., Osley,M.A., and Lipinski,M. (1995). A human homolog of the S. cerevisiae HIR1 and HIR2 transcriptional repressors cloned from the DiGeorge syndrome critical region. Hum. Mol. Genet. 4, 791-799.

Land,T. and Rouault,T.A. (1998). Targeting of a human iron-sulfur cluster assembly enzyme, nifs, to different subcellular compartments is regulated through alternative AUG utilization. Mol. Cell 2, 807-815.

Landino,L.M., Skreslet,T.E., and Alston,J.A. (2004). Cysteine oxidation of tau and microtubule-associated protein-2 by peroxynitrite: modulation of microtubule assembly kinetics by the thioredoxin reductase system. J. Biol Chem.

Latterich,M., Frohlich,K.U., and Schekman,R. (1995). Membrane fusion and the cell cycle: Cdc48p participates in the fusion of ER membranes. Cell 82, 885-893.

Lee,M.G. and Nurse,P. (1987). Complementation used to clone a human homologue of the fission yeast cell cycle control gene cdc2. Nature 327, 31-35.

Lippincott,J. and Li,R. (2000). Nuclear envelope fission is linked to cytokinesis in budding yeast. Exp. Cell Res. 260, 277-283.

Lockington,R., Scazzocchio,C., Sequeval,D., Mathieu,M., and Felenbok,B. (1987). Regulation of alcR, the positive regulatory gene of the ethanol utilization regulon of Aspergillus nidulans. Mol. Microbiol. 1, 275-281.

Lockington,R.A., Sealy-Lewis,H.M., Scazzocchio,C., and Davies,R.W. (1985). Cloning and characterization of the ethanol utilization regulon in Aspergillus nidulans. Gene 33, 137-149.

191 Lohka,M.J., Hayes,M.K., and Maller,J.L. (1988). Purification of maturation- promoting factor, an intracellular regulator of early mitotic events. Proc. Natl. Acad. Sci. U. S. A 85, 3009-3013.

Lorain,S., Lecluse,Y., Scamps,C., Mattei,M., and Lipinski,M. (2001). Identification of human and mouse HIRA-interacting protein-5 (HIRIP5), two mammalian representatives in a family of phylogenetically conserved proteins with a role in the biogenesis of Fe/S proteins. Biochim. Biophys. Acta 1517, 376- 383.

Lorain,S., Quivy,J.P., Monier-Gavelle,F., Scamps,C., Lecluse,Y., Almouzni,G., and Lipinski,M. (1998). Core histones and HIRIP3, a novel histone-binding protein, directly interact with WD repeat protein HIRA. Mol. Cell. Biol. 18, 5546-5556.

Lou,Y., Yao,J., Zereshki,A., Dou,Z., Ahmed,K., Wang,H., Hu,J., Wang,Y., and Yao,X. (2004). NEK2A interacts with MAD1 and possibly functions as a novel integrator of the spindle checkpoint signaling. J. Biol. Chem. 279, 20049-20057.

Lu,K.P. and Hunter,T. (1995a). Evidence for a NIMA-like mitotic pathway in vertebrate cells. Cell 81, 413-424.

Lu,K.P. and Hunter,T. (1995b). The NIMA kinase: A mitotic regulator in Aspergillus nidulans and vertebrate cells. Prog. Cell Cycle Res. 1, 187- 205.

Lu,K.P., Kemp,B.E., and Means,A.R. (1994). Identification of substrate specificity determinants for the cell cycle-regulated NIMA protein kinase. J. Biol. Chem. 269, 6603-6607.

Lu,K.P. and Means,A.R. (1994). Expression of the noncatalytic domain of the NIMA kinase causes a G2 arrest in Aspergillus nidulans. EMBO J. 13, 2103-2113.

Lu,K.P., Osmani,S.A., and Means,A.R. (1993). Properties and regulation of the cell cycle-specific NIMA protein kinase of Aspergillus nidulans. J. Biol. Chem. 268, 8769-8776.

Lupas,A., Van Dyke,M., and Stock,J. (1991). Predicting coiled coils from protein sequences. Science 252, 1162-1164.

Ma,J. and Ptashne,M. (1987). A new class of yeast transcriptional activators. Cell 51, 113-119.

192 Magnaghi,P., Roberts,C., Lorain,S., Lipinski,M., and Scambler,P.J. (1998). HIRA, a mammalian homologue of Saccharomyces cerevisiae transcriptional co- repressors, interacts with Pax3. Nat. Genet. 20, 74-77.

Marek,S.M., Wu,J., Louise,G.N., Gilchrist,D.G., and Bostock,R.M. (2003). Nuclear DNA degradation during heterokaryon incompatibility in Neurospora crassa. Fungal Genet. Biol. 40, 126-137.

Marshall,I.C.B. and Wilson,K.L. (1997). Nuclear envelope assembly after mitosis. Trends Cell Biol. 7, 69-74.

Martin-Lluesma,S., Stucke,V.M., and Nigg,E.A. (2002). Role of Hec1 in spindle checkpoint signaling and kinetochore recruitment of Mad1/Mad2. Science 297, 2267-2270.

Masui,Y. (1967). Relative roles of the pituitary, follicle cells, and progesterone in the induction of oocyte maturation in Rana pipiens. J. Exp. Zool. 166, 365- 375.

Masui,Y. and Markert,C.L. (1971). Cytoplasmic control of nuclear behavior during meiotic maturation of frog oocytes. J. Exp. Zool. 177, 129-145.

Mayor,T., Hacker,U., Stierhof,Y.D., and Nigg,E.A. (2002). The mechanism regulating the dissociation of the centrosomal protein C-Nap1 from mitotic spindle poles. J. Cell Sci. 115, 3275-3284.

Mayor,T., Stierhof,Y.D., Tanaka,K., Fry,A.M., and Nigg,E.A. (2000). The centrosomal protein C-Nap1 is required for cell cycle-regulated centrosome cohesion. J. Cell Biol. 151, 837-846.

Michaelis,C., Ciosk,R., and Nasmyth,K. (1997). Cohesins: chromosomal proteins that prevent premature separation of sister chromatids. Cell 91, 35-45.

Moreno,S., Hayles,J., and Nurse,P. (1989). Regulation of p34cdc2 protein kinase during mitosis. Cell 58, 361-372.

Moreno,S., Nurse,P., and Russell,P. (1990). Regulation of mitosis by cyclic accumulation of p80cdc25 mitotic inducer in fission yeast. Nature 344, 549- 552.

Morris,N.R. (1976). Mitotic mutants of Aspergillus nidulans. Genet. Res. Camb. 26, 237-254.

Mylyk,O.M. (1975). Heterokaryon incompatibility genes in Neurospora crassa detected using duplication-producing chromosome rearrangements. Genetics 80, 107-124.

193 Nakai,K. and Kanehisa,M. (1992). A knowledge base for predicting protein localization sites in eukaryotic cells. Genomics 14 , 897-911.

Nigg,E.A. (1998). Polo-like kinases: positive regulators of cell division from start to finish. Curr. Opin. Cell Biol. 10, 776-783.

Nikolay,R., Wiederkehr,T., Rist,W., Kramer,G., Mayer,M.P., and Bukau,B. (2004). Dimerization of the human E3 ligase CHIP via a coiled-coil domain is essential for its activity. J. Biol. Chem. 279, 2673-2678.

Nishitani,H. and Lygerou,Z. (2002). Control of DNA replication licensing in a cell cycle. Genes Cells 7, 523-534.

Nurse,P. (1994). Ordering S phase and M phase in the cell cycle. Cell 79, 547- 550.

Nurse,P. and Bisset,Y. (1981). Gene required in G1 for commitment to cell cycle and in G2 for control of mitosis in fission yeast. Nature 292, 558-560.

Nurse,P. and Thuriaux,P. (1980). Regulatory genes controlling mitosis in the fission yeast Schizosaccharomyces pombe. Genetics 96, 627-637.

Nurse,P., Thuriaux,P., and Nasmyth,K. (1976). Genetic control of the cell division cycle in the fission yeast Schizosaccharomyces pombe. Mol. Gen. Genet. 146, 167-178.

Nyberg,K.A., Michelson,R.J., Putnam,C.W., and Weinert,T.A. (2002). Toward maintaining the genome: DNA damage and replication checkpoints. Annu. Rev. Genet. 36, 617-656.

O'Connell,M.J., Krien,M.J., and Hunter,T. (2003). Never say never. the NIMA- related protein kinases in mitotic control. Trends Cell Biol. 13, 221-228.

O'Connell,M.J., Norbury,C., and Nurse,P. (1994). Premature chromatin condensation upon accumulation of NIMA. EMBO J. 13, 4926-4937.

Oakley,B.R. and Morris,N.R. (1983). A mutation in Aspergillus nidulans that blocks the transition from interphase to prophase. J. Cell Biol. 96, 1155- 1158.

Osmani,A.H., Davies,J., Oakley,C.E., Oakley,B.R., and Osmani,S.A. (2003). TINA interacts with the NIMA kinase in Aspergillus nidulans and negatively regulates astral microtubules during metaphase arrest. Mol. Biol. Cell 14, 3169-3179.

194 Osmani,A.H., McGuire,S.L., and Osmani,S.A. (1991a). Parallel activation of the NIMA and p34cdc2 cell cycle-regulated protein kinases is required to initiate mitosis in A. nidulans. Cell 67, 283-291.

Osmani,A.H., O'Donnell,K., Pu,R.T., and Osmani,S.A. (1991b). Activation of the nimA protein kinase plays a unique role during mitosis that cannot be bypassed by absence of the bimE checkpoint. EMBO J. 10, 2669-2679.

Osmani,S.A., Engle,D.B., Doonan,J.H., and Morris,N.R. (1988a). Spindle formation and chromatin condensation in cells blocked at interphase by mutation of a negative cell cycle control gene. Cell 52, 241-251.

Osmani,S.A., May,G.S., and Morris,N.R. (1987). Regulation of the mRNA levels of nimA, a gene required for the G2-M transition in Aspergillus nidulans. J. Cell Biol. 104, 1495-1504.

Osmani,S.A. and Mirabito,P.M. (2004). The early impact of genetics on our understanding of cell cycle regulation in Aspergillus nidulans. Fungal. Genet. Biol. 41, 401-410.

Osmani,S.A., Pu,R.T., and Morris,N.R. (1988b). Mitotic induction and maintenance by overexpression of a G2- specific gene that encodes a potential protein kinase. Cell 53, 237-244.

Osmani,S.A. and Ye,X.S. (1996). Cell cycle regulation in Aspergillus by two protein kinases. Biochem. J. 317, 633-641.

Perkins,D.D. (1988). Main features of vegetative incompatibility in Neurospora. Fungal Genetics Newsletter 35, 44-46.

Pitt,C.W., Moreau,E., Lunness,P.A., and Doonan,J.H. (2004). The pot1+ homologue in Aspergillus nidulans is required for ordering mitotic events. J. Cell Sci. 117, 199-209.

Pontecorvo,G. (1945). Biochemical and industrial use of moulds— application of genetical methods. Chem. Prod. 8, 41-43.

Pontecorvo,G. (1953). The genetics of Aspergillus nidulans. In Advances in Genetics, M.Demerec, ed. (New York: Academic Press), pp. 141-238.

Prigozhina,N.L., Oakley,C.E., Lewis,A.M., Nayak,T., Osmani,S.A., and Oakley,B.R. (2004). gamma-tubulin plays an essential role in the coordination of mitotic events. Mol. Biol. Cell 15, 1374-1386.

195 Pu,R.T., Gang Xu, Wu,L., Vierula,J., O'Donnell,K., Ye,X., and Osmani,S.A. (1995). Isolation of a functional homolog of the cell cycle specific NIMA protein kinase and functional analysis of conserved residues. J. Biol. Chem. 271, 18110-18116.

Pu,R.T. and Osmani,S.A. (1995). Mitotic destruction of the cell cycle regulated NIMA protein kinase of Aspergillus nidulans is required for mitotic exit. EMBO J. 14, 995-1003.

Rao,H., Uhlmann,F., Nasmyth,K., and Varshavsky,A. (2001). Degradation of a cohesin subunit by the N-end rule pathway is essential for chromosome stability. Nature 410, 955-959.

Rechsteiner,M. and Rogers,S.W. (1996). PEST sequences and regulation by proteolysis. Trends Biochem. Sci. 21, 267-271.

Robinow,C.F. and Caten,C.E. (1969). Mitosis in Aspergillus nidulans. J. Cell Sci. 5, 403-431.

Rogers,S., Wells,R., and Rechsteiner,M. (1986). Amino acid sequences common to rapidly degraded proteins: the PEST hypothesis. Science 234, 364-368.

Roig,J., Mikhailov,A., Belham,C., and Avruch,J. (2002). Nercc1, a mammalian NIMA-family kinase, binds the Ran GTPase and regulates mitotic progression. Genes Dev. 16, 1640-1658.

Rosenberger,R.F. and Kessel,M. (1967). Synchrony of nuclear replication in individual hyphae of Aspergillus nidulans. J. Bacteriol. 94, 1464-1469.

Rosenberger,R.F. and Kessel,M. (1968). Nonrandom sister chromatid segregation and nuclear migration in hyphae of Aspergillus nidulans. J. Bacteriol. 96, 1208-1213.

Roth,A.F. and Davis,N.G. (2000). Ubiquitination of the PEST-like endocytosis signal of the yeast a-factor receptor. J. Biol. Chem. 275, 8143-8153.

Rout,M.P. and Aitchison,J.D. (2001). The nuclear pore complex as a transport machine. J. Biol. Chem. 276, 16593-16596.

Russell,P. and Nurse,P. (1986). cdc25 functions as an inducer in the mitotic control of fission yeast. Cell 45, 145-153.

Russell,P. and Nurse,P. (1987). The mitotic inducer nim1+ functions in a regulatory network of protein kinase homologs controlling the initiation of mitosis. Cell 49, 569-576.

196 Ryan,K.J., McCaffery,J.M., and Wente,S.R. (2003). The Ran GTPase cycle is required for yeast nuclear pore complex assembly. J. Cell Biol. 160, 1041- 1053.

Salina,D., Bodoor,K., Eckley,D.M., Schroer,T.A., Rattner,J.B., and Burke,B. (2002). Cytoplasmic dynein as a facilitator of nuclear envelope breakdown. Cell 108, 97-107.

Sarkar,S., Iyer,G., Wu,J., and Glass,N.L. (2002). Nonself recognition is mediated by HET-C heterocomplex formation during vegetative incompatibility. EMBO J. 21, 4841-4850.

Saupe,S.J., Clave,C., Sabourin,M., and Begueret,J. (2000). Characterization of hch, the Podospora anserina homolog of the het-c heterokaryon incompatibility gene of Neurospora crassa. Curr. Genet. 38, 39-47.

Saupe,S.J. and Glass,N.L. (1997). Allelic specificity at the het-c heterokaryon incompatibility locus of Neurospora crassa is determined by a highly variable domain. Genetics 146, 1299-1309.

Saupe,S.J., Kuldau,G.A., Smith,M.L., and Glass,N.L. (1996). The product of the het-C heterokaryon incompatibility gene of Neurospora crassa has characteristics of a glycine-rich cell wall protein. Genetics 143, 1589-1600.

Sauve,D.M., Anderson,H.J., Ray,J.M., James,W.M., and Roberge,M. (1999). Phosphorylation-induced rearrangement of the histone H3 NH2-terminal domain during mitotic chromosome condensation. J. Cell. Biol. 145, 225- 235.

Schilke,B., Voisine,C., Beinert,H., and Craig,E. (1999). Evidence for a conserved system for iron metabolism in the mitochondria of Saccaromyces cerevisiae. Proc. Natl. Acad. Sci. U. S. A. 96, 10206-10211.

Schultz,L.B., Chehab,N.H., Malikzay,A., DiTullio,R.A., Jr., Stavridi,E.S., and Halazonetis,T.D. (2000). The DNA damage checkpoint and human cancer. Cold Spring Harb. Symp. Quant. Biol. 65, 489-498.

Simanis,V. and Nurse,P. (1986). The cell cycle control gene cdc2+ of fission yeast encodes a protein kinase potentially regulated by phosphorylation. Cell 45, 261-268.

Steinberg,R.A., Cauthron,R.D., Symcox,M.M., and Shuntoh,H. (1993). Autoactivation of catalytic (Ca) subunit of cyclic AMP-dependent protein kinase by phosphorylation of threonine 197. Mol. Cell. Biol. 13, 2332-2341.

Stemmann,O., Zou,H., Gerber,S.A., Gygi,S.P., and Kirschner,M.W. (2001). Dual inhibition of sister chromatid separation at metaphase. Cell 107, 715-726. 197 Sveiczer,A., Tyson,J.J., and Novak,B. (2004). Modelling the fission yeast cell cycle. Brief. Funct. Genomic. Proteomic. 2, 298-307.

Swaminathan,S., Ellis,H.M., Waters,L.S., Yu,D., Lee,E.C., Court DL, and Sharan,S.K. (2001). Rapid engineering of bacterial artificial chromosomes using oligonucleotides. Genesis. 29, 14-21.

Swenson,K.I., Farrell,K.M., and Ruderman,J.V. (1986). The clam embryo protein cyclin A induces entry into M phase and the resumption of meiosis in Xenopus oocytes. Cell 47, 861-870.

Tanaka,T., Cosma,M.P., Wirth,K., and Nasmyth,K. (1999). Identification of cohesin association sites at centromeres and along chromosome arms. Cell 98, 847-858.

Tanaka,T., Fuchs,J., Loidl,J., and Nasmyth,K. (2000). Cohesin ensures bipolar attachment of microtubules to sister centromeres and resists their precocious separation. Nat. Cell Biol. 2, 492-499.

Taylor,S.S., Buechler,J.A., and Yonemoto,W. (1990). cAMP-dependent protein kinase: framework for a diverse family of regulatory . Annu. Rev. Biochem. 59, 971-1005.

Todd,N.K. and Rayner,A.D. (1980). Fungal individualism. Sci. Prog. Oxford 66, 331-354.

Tomonaga,T., Nagao,K., Kawasaki,Y., Furuya,K., Murakami,A., Morishita,J., Yuasa,T., Sutani,T., Kearsey,S.E., Uhlmann,F., Nasmyth,K., and Yanagida,M. (2000). Characterization of fission yeast cohesin: essential anaphase proteolysis of Rad21 phosphorylated in the S phase. Genes Dev. 14 , 2757-2770.

Tong,W.H., Jameson,G.N., Huynh,B.H., and Rouault,T.A. (2003). Subcellular compartmentalization of human Nfu, an iron-sulfur cluster scaffold protein, and its ability to assemble a [4Fe-4S] cluster. Proc. Natl. Acad. Sci. U. S. A. 100, 9762-9767.

Toth,A., Ciosk,R., Uhlmann,F., Galova,M., Schleiffer,A., and Nasmyth,K. (1999). Yeast cohesin complex requires a conserved protein, Eco1p(Ctf7), to establish cohesion between sister chromatids during DNA replication. Genes Dev. 13, 320-333.

Uhlmann,F., Lottspeich,F., and Nasmyth,K. (1999). Sister-chromatid separation at anaphase onset is promoted by cleavage of the cohesin subunit Scc1. Nature 400, 37-42.

198 Uhlmann,F. and Nasmyth,K. (1998). Cohesion between sister chromatids must be established during DNA replication. Curr. Biol. 8, 1095-1101.

Uhlmann,F., Wernic,D., Poupart,M.A., Koonin,E.V., and Nasmyth,K. (2000). Cleavage of cohesin by the CD clan protease separin triggers anaphase in yeast. Cell 103, 375-386.

Uto,K., Nakajo,N., and Sagata,N. (1999). Two structural variants of Nek2 kinase, termed Nek2A and Nek2B, are differentially expressed in Xenopus tissues and development. Dev. Biol. 208, 456-464.

Van Hooser,A., Goodrich,D.W., Allis,C.D., Brinkley,B.R., and Mancini,M.A. (1998). Histone H3 phosphorylation is required for the initiation, but not maintenance, of mammalian chromosome condensation. J. Cell. Sci. 111, 3497-3506.

Walther,T.C., Alves,A., Pickersgill,H., Loiodice,I., Hetzer,M., Galy,V., Hulsmann,B.B., Kocher,T., Wilm,M., Allen,T., Mattaj,I.W., and Doye,V. (2003a). The conserved Nup107-160 complex is critical for nuclear pore complex assembly. Cell 113, 195-206.

Walther,T.C., Askjaer,P., Gentzel,M., Habermann,A., Griffiths,G., Wilm,M., Mattaj,I.W., and Hetzer,M. (2003b). RanGTP mediates nuclear pore complex assembly. Nature 424, 689-694.

Waring,R.B., May,G.S., and Morris,N.R. (1989). Characterization of an inducible expression system in Aspergillus nidulans using alcA and tubulin-coding genes. Gene 79, 119-130.

Wei,Y., Yu,L.L., Bowen,J., Gorovsky,M.A., and Allis,C.D. (1999). Phosphorylation of histone H3 is required for proper chromosome condensation and segregation. Cell 97, 99-109.

Wente,S.R. (2000). Gatekeepers of the nucleus. Science 288, 1374-1377.

Wolkow,T.D., Harris,S.D., and Hamer,J.E. (1996). Cytokinesis in Aspergillus nidulans is controlled by cell size, nuclear positioning and mitosis. J. Cell Sci. 109, 2179-2188.

Wosten,H.A., Moukha,S.M., Sietsma,J.H., and Wessels,J.G. (1991). Localization of growth and secretion of proteins in Aspergillus niger. J. Gen. Microbiol. 137, 2017-2023.

Wozniak,R. and Clarke,P.R. (2003). Nuclear pores: sowing the seeds of assembly on the chromatin landscape. Curr. Biol. 13, R970-R972.

199 Wu,J. and Glass,N.L. (2001). Identification of specificity determinants and generation of alleles with novel specificity at the het-c heterokaryon incompatibility locus of Neurospora crassa. Mol. Cell. Biol. 21, 1045-1057.

Wu,L., Osmani,S.A., and Mirabito,P.M. (1998). A role for NIMA in the nuclear localization of cyclin B in Aspergillus nidulans. J. Cell Biol. 141, 1575-1587.

Yaglom,J., Linskens,M.H.K., Sadis,S., Rubin,D.M., Futcher,B., and Finley,D. (1995). p34Cdc28-mediated control of Cln3 cyclin degredation. Mol. Cell. Biol. 15, 731-741.

Ye,X.S., Fincher,R.R., Tang,A., O'Donnell,K., and Osmani,S.A. (1996). Two S- phase checkpoint systems, one involving the function of both BIME and Tyr15 phosphorylation of p34cdc2, inhibit NIMA and prevent premature mitosis. EMBO J. 15, 3599-3610.

Ye,X.S., Fincher,R.R., Tang,A., Osmani,A.H., and Osmani,S.A. (1998). Regulation of the anaphase-promoting complex/cyclosome by BIMA (APC3) and proteolysis of NIMA. Mol. Biol. Cell 9, 3019-3030.

Ye,X.S., Xu,G., Pu,P.T., Fincher,R.R., McGuire,S.L., Osmani,A.H., and Osmani,S.A. (1995). The NIMA protein kinase is hyperphosphorylated and activated downstream of p34cdc2/cyclin B: coordination of two mitosis promoting kinases. EMBO J. 14, 986-994.

Yin,M.J., Shao,L., Voehringer,D., Smeal,T., and Jallal,B. (2003). The serine/threonine kinase Nek6 is required for cell cycle progression through mitosis. J. Biol. Chem. 278, 52454-52460.

Yu,D., Ellis,H.M., Lee,E.C., Jenkins,N.A., Copeland,N.G., and Court DL (2000). An efficient recombination system for chromosome engineering in Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 97, 5978-5983.

Yu,H. (2002). Regulation of APC-Cdc20 by the spindle checkpoint. Curr. Opin. Cell Biol. 14, 706-714.

Yu,H., Peters,J.M., King,R.W., Page,A.M., Hieter,P., and Kirschner,M.W. (1998). Identification of a cullin homology region in a subunit of the anaphase- promoting complex. Science 279, 1219-1222.

Yuvaniyama,P., Agar,J.N., Cash,V.L., Johnson,M.K., and Dean,D.R. (2000). NifS-directed assembly of a transient [2Fe-2S] cluster within the NifU protein. Proc. Natl. Acad. Sci. U. S. A. 97, 599-604.

Zheng,L., Chen,Y., and Lee,W.H. (1999). Hec1p, an evolutionarily conserved coiled-coil protein, modulates chromosome segregation through interaction with SMC proteins. Mol. Cell. Biol. 19, 5417-5428. 200 Zou,H., McGarry,T.J., Bernal,T., and Kirschner,M.W. (1999). Identification of a vertebrate sister-chromatid separation inhibitor involved in transformation and tumorigenesis. Science 285, 418-422.

Zou,H., Stemman,O., Anderson,J.S., Mann,M., and Kirschner,M.W. (2002). Anaphase specific auto-cleavage of separase. FEBS Lett. 528, 246-250.

201