Anillin Stabilizes Membrane-Cytoskeleton Interactions During Drosophila Male Germ Cell Cytokinesis

Philip Daniel Goldbach

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy The Department of Molecular Genetics University of Toronto

Philip Daniel Goldbach

Doctor of Philosophy

The Department of Molecular Genetics University of Toronto

2011 Abstract

The scaffolding protein anillin plays a crucial role during cytokinesis – the physical separation of daughter cells following chromosome segregation. Anillin binds filamentous F-actin, non- muscle myosin II and septins, and in cell culture models has been shown to restrict actomyosin contractility to the cleavage furrow. Whether anillin also serves this function during the incomplete cytokinesis that occurs in developing germ cells has remained unclear. Localization of anillin to several actin-rich structures in developing male germ cells also suggests potential roles for anillin outside of cytokinesis. In this study, I demonstrate that anillin is required for cytokinesis in dividing Drosophila spermatocytes. In addition, spermatid individualization is defective in anillin-depleted cells, although similarities to another cytokinesis mutant, four wheel drive, suggest this may be a secondary effect of failed cytokinesis. Anillin, septins and myosin II stably associate with the cleavage furrow in wild-type dividing spermatocytes. Anillin is necessary for recruitment of septins to the cleavage furrow, and for maintenance of Rho, F-actin and myosin II at the equator in late stages of cytokinesis. Membrane trafficking appears unaffected in anillin-depleted cells, although, unexpectedly, ectopic expression of one membrane trafficking marker, DE-cadherin-GFP, suppresses the cytokinesis defect. DE-cadherin-GFP recruits β-catenin (armadillo) and α-catenin to the cleavage furrow and stabilizes F-actin at the ii equator. Taken together, my results suggest that the anillin-septin and cadherin-catenin complexes can serve as alternative means to promote tight physical coupling of F-actin and myosin II to the cleavage furrow and successful completion of cytokinesis.

iii

Acknowledgments

A number of people must be thanked for their help and guidance over the past several years. First, I would like to express my sincere gratitude to and appreciation for my supervisor, Julie Brill. Throughout my time in the lab, she has been supportive and exceedingly generous with her time. Her dedication, depth of knowledge and scientific curiosity have been truly inspiring. In addition to making me a better scientist, she has made me a better writer and editor (and thinker). She’s also shown me how one can succeed in the scientific community as a collaborator rather than a competitor. Whatever the future holds, the skills I’ve developed in Julie’s lab will undoubtedly serve me well.

I would also like to thank my committee members, Andrew Wilde and Howard Lipshitz. They’ve been very helpful in keeping me focused and on track. I remain deeply impressed by their sharpness and keen eyes for good, solid science.

Of course, I couldn’t write this without acknowledging my wonderful, and sometimes bizarre, labmates. Ho-Chun, Gordon, Jason and Julie T. have been with me through most of the journey and have been never-ending sources of technical and moral support (and, of course, mockery). There have been plenty of other people in the lab who have been generous with their time and expertise and I offer you all my gratitude. Overall, as inevitably happens, the lab becomes a home away from home and the members part of an extended family. It’s a unique home (and family) and I’ll definitely miss it.

I would like to thank a number of people for kindly providing expertise and/or reagents. Crucially, our collaborators, Nolan Beise and Bill Trimble, for carrying out the L-fibroblast experiments and contributing to the anillin story; Angela Barth and James Nelson for providing them with reagents; and Ritu Sarpal for providing the alpha-catenin antibody. I would also like to thank Zhigang Jin and Shelagh Campbell for collaborations on the dMyt1 project. Additionally, Henry Chang, Karen Hales, Roger Karess, Hiroki Oda, Ulrich Tepass and the Bloomington Drosophila stock center for fly stocks; William Bement, Brian Burkel, Helmut Krämer, Erik Snapp, Roger Tsien and the Canadian Drosophila Microarray Centre for plasmids; Paul Paroutis and the Sick Kids Imaging Facility for help with FRAP experiments; and the folks

iv at Sick Kids Lab Animal Services, with whom I worked closely in generating the anillin antibody.

Last but not least, I would like to thank my amazing friends and family who have supported me throughout. Special mentions go to fellow Scooby Gang members and to all the roommates who’ve put up with me at 100 Spadina. I’ve been blessed with an extremely close immediate and extended family and their support is invaluable. Above all, I would like to thank my incredible parents, Morty and Elaine. Thank you.

Table of Contents

Acknowledgments...... iv

Table of Contents...... vi

List of Tables ...... ix

List of Figures ...... x

List of Abbreviations ...... xii

1 Introduction ...... 2

1.1 Cytokinesis...... 2

1.1.1 The stages of cytokinesis ...... 3

1.1.2 The main players: proteins and lipids that regulate, organize and guide cytokinesis...... 7

1.1.3 Membrane-cytoskeletal attachments in cytokinesis...... 15

1.1.4 The current cytokinesis picture: complexity, redundancy and open questions..... 17

1.2 Anillin and anillin-related proteins ...... 17

1.2.1 Anillin structure ...... 18

1.2.2 Anillin expression and localization...... 21

1.2.3 Known and predicted functions of anillin...... 22

1.3 The system: Drosophila spermatogenesis ...... 25

1.3.1 Overview...... 25

1.3.2 Meiotic cytokinesis ...... 28

1.3.3 Individualization ...... 32

1.4 Thesis rationale and summary...... 32

2 Materials and Methods...... 36

2.1 Molecular biology...... 36

2.2 Generation of anti-anillin antibody...... 37

2.3 Drosophila genetics ...... 39 vi

2.4 Fluorescence microscopy, imaging and analysis ...... 40

2.5 FRAP image acquisition and data analysis...... 41

2.6 Immunoblotting...... 41

3 Results ...... 44

3.1 Introduction...... 44

3.2 Results...... 46

3.2.1 Anillin is found in a variety of actin-associated structures during spermatogenesis ...... 46

3.2.2 Anillin is required for spermatocyte cytokinesis ...... 51

3.2.3 Anillin knockdown causes defects in investment cone morphology and function ...... 55

3.2.4 Anillin, septins and myosin II are stably associated with the contractile ring...... 57

3.2.5 Anillin recruits septins and maintains Rho, F-actin and myosin II at the cleavage furrow...... 59

3.2.6 Ectopically expressed DE-cad-GFP as a marker for membrane trafficking in male germ cells ...... 64

3.2.7 Loss of anillin is suppressed by expression of DE-cadherin ...... 64

3.2.8 DE-cad-GFP recruits armadillo and α-catenin to the cleavage furrow ...... 68

3.2.9 Adherens junction proteins can substitute for anillin in anchoring F-actin at the cleavage furrow...... 71

3.2.10 The ability of E-cadherin to suppress loss of anillin is conserved in mouse L- fibroblasts...... 74

3.3 Conclusions...... 76

4 Discussion and Future Directions ...... 79

4.1 Anillin in contractile ring-membrane attachment ...... 79

4.1.1 Dynamics of contractile ring components during cytokinesis...... 79

4.1.2 Anillin as a cytoskeletal-membrane crosslinker ...... 81

4.1.3 Proposal: Testing phospholipid-binding ability of anillin ...... 82

4.2 Anillin function(s) in Drosophila spermatogenesis...... 84 vii

4.2.1 Individualization ...... 85

4.2.2 Anillin and septins in ring canal stability ...... 85

4.2.3 Anillin stability and regulation ...... 86

4.2.4 Proposal: Examining post-meiotic functions of anillin ...... 87

4.3 A role for E-cadherin in animal cell cytokinesis?...... 89

4.3.1 Mechanism of E-cadherin suppression ...... 89

4.3.2 An endogenous role for E-cadherin in cytokinesis? ...... 91

4.3.3 Proposal: Determining the mechanism of E-cadherin suppression ...... 93

4.4 Concluding remarks ...... 94

Appendix...... 96

References...... 110

viii

List of Tables

Table 1.1 Genes with roles in male meiotic cytokinesis...... 31

List of Figures

Figure 1.1. Models of Cytokinesis...... 5

Figure 1.2. Schematic representation of the Drosophila anillin protein...... 19

Figure 1.3. Overview of Drosophila spermatogenesis and spermatid individualization...... 27

Figure 1.4. Drosophila male meiotic cytokinesis...... 28

Figure 3.1. Generation of anti-anillin antibody...... 48

Figure 3.2. Anillin localizes to actin-rich structures during spermatogenesis...... 49

Figure 3.3. GFP- and RFP-anillin localize to same structures recognized by anti-anillin antibody...... 50

Figure 3.4. Anillin is required for spermatocyte cytokinesis...... 52

Figure 3.5. Anillin is required for normal investment cone morphology...... 56

Figure 3.6. Anillin, septins and myosin II are stably associated with the cleavage furrow...... 58

Figure 3.7. Anillin is required for recruitment of septins and stabilization of Rho and myosin II at the cleavage furrow...... 61

Figure 3.8. Anillin is required for tight association of myosin II with the cleavage furrow...... 63

Figure 3.9. DE-cad-GFP suppresses cytokinesis defects caused by anillin depletion...... 66

Figure 3.10. Membrane trafficking appears unaffected in anillin-depleted cells...... 67

Figure 3.11. Endogenous DE-cadherin, armadillo and α-catenin are not detected in dividing spermatocytes...... 69

Figure 3.12. DE-cad-GFP recruits armadillo and α-catenin to the cleavage furrow...... 70

Figure 3.13. Anillin and DE-cadherin restrict F-actin to the cleavage furrow...... 72

Figure 3.14. DE-cadherin expression does not rescue peanut localization in anillin-depleted cells...... 73

Figure 3.15. Expression of E-cadherin suppresses cytokinesis defects caused by depletion of anillin in mouse L cells...... 75

Figure 4.1. Cadherin-catenins can replace anillin-septins in stabilizing the furrow during cytokinesis...... 91

List of Abbreviations

α-cat: alpha-catenin

AHR: anillin homology region

Anil: anillin

APC/C: anaphase promoting complex/cyclosome

Arm: armadillo, Drosophila beta-catenin

ATP: adenosine triphosphate

ATPase: adenosine triphosphatase

β2t: beta 2-tubulin

C-terminus: carboxy terminus

Cdk1: cyclin dependent kinase 1 cDNA: complementary DNA

Chic: chickadee, a Drosophila profilin

CNS: central nervous system

CR: contractile ring

DE-cad: Drosophila E-cadherin

Dia: diaphanous, a Drosophila formin dsRNA: double stranded RNA

E-cad: E-cadherin

ERM: ezrin/radixin/moesin

ESCRT: endosomal sorting complex required for transport

F-actin: filamentous actin

FRAP: fluorescence recovery after photobleaching

Fwd: four wheel drive, Drosophila phosphatidylinositol 4-kinase III beta xii

GAP: GTPase activating protein

GEF: GDP-GTP exchange factor

GFP: green fluorescent protein

GTP: guanosine triphosphate

GTPase: guanosine triphosphatase

IQGAP: IQ motif containing GTPase activating protein mDia: mammalian diaphanous-related, a formin

MgcRacGAP: male germ cell Rac-GTPase-activating protein

MKLP1: mitotic kinesin-like protein 1

MLCK: myosin light chain kinase

MYPT: myosin phosphatase

N-terminus: amino terminus

NES: nuclear exit signal

NLS: nuclear localization signal

Pav: pavarotti, a Drosophila kinesin-like protein

Pbl: pebble, Drosophila RhoGEF

PCH: pombe Cdc15 homology

PE: phosphatidylethanolamine

Pex: peroxin

PH domain: pleckstrin homology domain

PI: phosphatidylinositol

PI3K: phosphoinositide 3-kinase

PI4K: phosphatidylinositol 4-kinase

PI4P: phosphatidylinositol 4-phosphate

PI5P: phosphatidylinositol 5-phosphate

PIP: phosphatidylinositol phosphate xiii

PIP2: phosphatidylinositol 4,5-bisphosphate

PIP3: phosphatidylinositol 3,4,5-triphosphate

PIP5K: phosphatidylinositol 4-phosphate 5-kinase

PLCδ: phospholipase C-delta

PLK1: polo-like kinase 1

PNS: peripheral nervous system

Pnut: peanut, a septin

RacGAP: Rac GTPase-activating protein

RFP: red fluorescent protein

RNAi: RNA interference

ROCK: Rho-dependent kinase scra: scraps, original Drosophila name of anillin-encoding gene

SDS-PAGE: sodium dodecyl sulphate polyacrylamide gel electrophoresis sGFP: secreted green fluorescent protein

SH3 domain: src homology-3 domain shRNA: short hairpin RNA

Sktl: skittles, a Drosophila PIP 5-kinase

SNARE: soluble N-ethylmaleimide-sensitive factor activating protein receptor

Sqh: spaghetti squash, Drosophila myosin regulatory light chain

TCE: translational controlled element

TEV: tobacco etch virus

Tsr: twinstar, a Drosophila cofilin

UAS: upstream activating sequence

UTR: untranslated region

Utr-CH: utrophin calponin-homology domain

VDRC: Vienna Drosophila RNAi Collection xiv 1

Chapter 1

Introduction

1 Introduction 1.1 Cytokinesis

Cytokinesis – the physical separation of two daughter cells following the segregation of genetic materials – was initially described well over a hundred years ago by biologists examining living cells using the latest microscope technology (Rappaport, 1996). Many theories were posited shortly thereafter to describe the mechanism of division. These often drew on analogies to physical and mechanical properties of inorganic materials, and relied on scant experimental evidence. Little was known about the molecular makeup of cells in general and the cytokinetic apparatus in particular. As a result little progress in understanding cytokinesis was made over a long period of time. This has changed over the last twenty years with several important advances. One is the assembly of lists of specific molecules that contribute to cytokinesis in different organisms (Glotzer, 2005; Eggert et al., 2006). Another is the advance in technology, which, crucially, has allowed researchers to a) manipulate cells in more and more precise ways and b) generate computational models to test specific theories of cytokinesis.

It was once assumed that cytokinesis was a relatively simple mechanical process that could be understood by uncovering certain principles that apply to all dividing cells. Such is no longer the case. Instead, many reports, often in conflict with each other, have revealed cytokinesis to be a complex process with redundant and overlapping systems, the importance of which varies in different cell types and under different conditions. Even with this complexity, there appears to be a core set of molecules and general properties shared by most cells. The data presented in this thesis contribute to this emerging picture, revealing general as well as specific features of the scaffolding protein, anillin, and of the cytokinetic apparatus.

In the following section, I will provide a summary of the current state of knowledge in the cytokinesis field. The focus will be on cytokinesis in animal cells and in the fission yeast, Schizosaccharomyces pombe. Descriptions of cytokinesis in plants, bacteria and other types of yeast are, for the large part, outside the scope of this thesis. I will begin with a description of the basic stages of cytokinesis.

1.1.1 The stages of cytokinesis

Cytokinesis can be subdivided into specific stages. Although not discussed here, tight coordination of chromosome segregation and cytokinesis is crucial to prevent errors during division. The first stage of cytokinesis occurs during anaphase with specification of the division site.

1.1.1.1 Division site specification and contractile ring formation

A long-standing debate in the cytokinesis field deals with the nature and origin of the signal that determines where along the plasma membrane division occurs. In most cell types, division occurs at the equator of the cell, equidistant from the spindle poles. There are plenty of examples of cells, however, that undergo asymmetric division, indicating that, as in other aspects of cytokinesis, there is room for variation. Because of the clear relationship between the location and orientation of the mitotic spindle and the division site, the spindle was long seen as a potential source of activating and/or inhibitory signals involved in setting up the division site. The mitotic spindle can be subdivided into distinct populations of microtubules: astral microtubules that radiate from centrosomes to the cell periphery; kinetochore microtubules (also called chromosomal microtubules) that link chromosomes to centrosomes; and the central spindle (also called interzonal or midzone microtubules) – a group of interdigitating microtubules that become compressed during furrow ingression and are later incorporated into the midbody (Figure 1.1A) (see Section 1.1.1.3). Which subset of microtubules provides the signal to divide remains an ongoing source of investigation. At one extreme are Caenorhabditis elegans embryos mutant for spd-1 that can divide in the complete absence of a central spindle (Verbrugghe and White, 2004); at the other extreme are Drosophila melanogaster spermatocytes mutant for asterless that can divide in the complete absence of astral microtubules (Bonaccorsi et al., 1998). Between these extremes are cells that use different microtubule populations as sources of complementary or overlapping signals (for recent reviews see Burgess and Chang, 2005; D'Avino et al., 2005; von Dassow, 2009). Regardless of the exact source, it is clear that some part of the mitotic spindle is essential for specifying the division plane in animal cells.

Another longstanding and ongoing debate deals with the nature of forces driving division. The most common form of this debate is between the concepts of equatorial stimulation and polar relaxation, both of which, in theory, could result in furrow formation and ingression at the

4 centre of the cell (Figure 1.1B) (Rappaport, 1996). The equatorial stimulation model posits that signals delivered to the equatorial cortex, as discussed above, initiate positive and localized inward forces. In contrast, the polar relaxation model posits that inhibitory signals delivered to the poles locally inhibit cortical tension; as a result, tension becomes highest at the equator, which leads to formation of a cleavage furrow. These mechanisms are not mutually exclusive. In many cell types, both equatorial stimulation and polar relaxation may contribute to varying degrees under different conditions (Rappaport, 1996; Burgess and Chang, 2005; D'Avino et al., 2005).

A molecular explanation for force generation in dividing cells was eventually suggested from key findings in the 1960s and '70s that the cleavage furrow contains filaments composed of filamentous actin (F-actin) and non-muscle myosin II (Schroeder, 1968; Forer and Behnke, 1972; Schroeder, 1973; Fujiwara et al., 1978). It is now generally accepted that a contractile ring (CR) of F-actin and myosin II provide most, if not all, of the force required for ingression of the cleavage furrow in animal cells (Robinson and Spudich, 2004). As in other aspects of cytokinesis, exceptions can be found. For example, the slime mold Dictyostelium discoideum can use polar traction forces to separate daughter cells on solid substrates, even in the absence of myosin II (Neujahr et al., 1997).

Similar to most metazoan cells, S. pombe assembles an actomyosin ring at the site of division. However, the method of division site selection differs markedly. Multiple experiments have demonstrated that for S. pombe the location of the nucleus, rather than the spindle, plays the key role (Burgess and Chang, 2005). When the nucleus is displaced by experimental manipulation, the division site relocates to the part of the membrane directly overlying the new nuclear position (Chang et al., 1996; Tran et al., 2001; Daga and Chang, 2005). Microtubules still play a role, however, in determining the location of the nucleus itself (Toda et al., 1983; Tran et al., 2001).

In summary, specific subpopulations of microtubules or nuclear positioning, in the case of S. pombe, are responsible for delivery of molecular signals to the cell cortex at the presumptive site of division. This in turn leads to localization, polymerization and activation of an actomyosin ring. A list of conserved molecules involved in these and later cytokinetic processes are discussed in more detail in Section 1.1.2.

Figure 1.1. Models of Cytokinesis.

(A,B) Schematic diagrams of animal cells undergoing conventional cell division. (A) The presumptive site of division (red dotted line) is selected by microtubules during anaphase (left cell). Astral microtubules (black lines) and microtubules of the central spindle (overlapping blue lines) contribute to site selection in different cell types. After cleavage site selection (right cell), a contractile ring of F-actin and myosin II (red), in association with additional proteins such as anillin and septins, constricts, leading to formation of a cleavage furrow. Interactions between the contractile ring and central spindle are important for cytokinesis completion. (B) Two longstanding theories offer alternative explanations for physical constriction of membranes between spindle poles. Equatorial stimulation (left cell) posits that microtubules deliver a positive cue (+) to the equatorial cortex, inducing inward force. In contrast, polar relaxation (right cell) posits that a tension gradient is created such that tension is lowest (−) at the cell poles. Higher relative tension at the equator results in localized constriction. These theories are not mutually exclusive and both may contribute to cleavage furrow formation.

1.1.1.2 Cleavage furrow ingression

Once actin and myosin filaments have been assembled at the site of division, and myosin is activated through phosphorylation of its regulatory light chain, the CR begins to contract. The contractile ring hypothesis states that contractile filaments are drawn together in a “purse-string”- like manner by myosin II activity (Satterwhite and Pollard, 1992; Wang, 2005). As a result, the membrane overlying the CR begins to pinch inward and a cleavage furrow becomes visible. As will be discussed in greater detail below, an intimate link must be formed and maintained between the CR and the membrane as the furrow ingresses.

One ongoing area of investigation focuses on CR component dynamics during cleavage furrow ingression. Because the ring shrinks in diameter as it constricts, it must be assumed that filaments are removed or disassembled during this process. This is in contrast to muscle fibers, which maintain stable configurations of actin and myosin in sarcomeres during activation. Recent studies making use of fluorescence recovery after photobleaching (FRAP) in yeast and mammalian cells suggest that F-actin and myosin are highly dynamic during constriction, leading to an idea that F-actin and myosin are constantly recruited and disassembled at the cleavage furrow during cytokinesis (Pelham and Chang, 2002; Murthy and Wadsworth, 2005). However, similar experiments in the early C. elegans embryo, published concurrent with experiments described in this thesis, suggest that actin and myosin do not turn over, but rather disassemble over time (Carvalho et al., 2009).

1.1.1.3 Completion of cytokinesis

As cytokinesis proceeds, the cleavage furrow ingresses until it is blocked by tightly bundled midbody microtubules. Completion of cytokinesis involves the coordination of multiple processes, including membrane trafficking and protein degradation, and culminates in the severing of the remaining connection between daughter cells by a process termed abscission (Steigemann and Gerlich, 2009). Many studies over the last decade have clearly established membrane trafficking as an important factor in later stages of cytokinesis (reviewed in Albertson et al., 2005; Barr and Gruneberg, 2007). The functions of trafficked membranes and associated proteins, in addition to the relative importance of different trafficking pathways such as endocytosis, recycling and secretion, remain open questions. Other requirements for proper completion of cytokinesis include disassembly of the CR and removal or inactivation of

7 regulatory molecules. Several proteins with roles in cytokinesis, including polo-like kinase 1 (PLK1) and anillin, are ubiquitinated by the anaphase promoting complex/cyclosome (APC/C), leading to proteolytic degradation (Lindon and Pines, 2004; Zhao and Fang, 2005). The final event is the severing of the midbody bridge, the timing of which varies greatly across cell types. Membrane fission mediated by endosomal sorting complex required for transport (ESCRT) is required for this step in mammalian cells (Carlton and Martin-Serrano, 2007; Morita et al., 2007; Carlton et al., 2008; Lee et al., 2008). Other proposed mechanisms for abscission include mechanical rupture and filling in of the intercellular space by internal vesicles (Steigemann and Gerlich, 2009). These processes are not mutually exclusive and may be more or less important under different conditions.

In summary, cytokinesis involves multiple steps or processes beginning with specification of the division plane and ending with abscission. Each step requires a number of conserved proteins and lipids, which I will describe in the following section. Special attention is given to those that interact with, or are otherwise important for the function of, the scaffolding protein anillin.

1.1.2 The main players: proteins and lipids that regulate, organize and guide cytokinesis

In the past several years, functions for specific proteins in cytokinesis have been discovered through classical genetics and biochemistry (reviewed in Glotzer, 2005; Eggert et al., 2006). In addition, large-scale screens making use of RNA interference (RNAi) technology in Drosophila, C. elegans and mammalian cells have generated lists of proteins involved in the process (Gonczy et al., 2000; Somma et al., 2002; Kiger et al., 2003; Echard et al., 2004; Eggert et al., 2004; Kittler et al., 2004; Zhu et al., 2005; Kittler et al., 2007). Although unique proteins were uncovered in different screens, it is clear that a core group of molecules is required for cytokinesis across a wide variety of cell types and organisms. This includes cytoskeletal and cytoskeleton-associated proteins, septins, membrane lipids, and other regulatory proteins. Anillin and anillin-related proteins, which are part of the core cytokinetic machinery, will be discussed separately in Section 1.2.

1.1.2.1 F-actin, non-muscle myosin II and associated proteins

As described above, the essential force-generating apparatus in almost all animal and fungal dividing cells is a CR composed of actin and myosin filaments. The details of how these proteins get to the division site and whether they arrive as monomers or previously formed filaments are still being uncovered. Actin filament assembly during cytokinesis requires actin nucleating formins, mDia2 in mammals and Diaphanous (Dia) in Drosophila, plus the actin monomer binding protein profilin, Chickadee (Chic) in Drosophila. Formins associate with the fast-growing barbed ends of actin filaments and control processive addition of profilin-bound actin monomers (Pruyne et al., 2002; Zigmond et al., 2003; Kovar and Pollard, 2004). The Arp2/3 complex, which nucleates branched actin filaments, has also been identified in multiple screens, although its role in cytokinesis is not as well established as that of formins (Eggert et al., 2004; Skop et al., 2004). Actin filaments are crosslinked in the area of the furrow and one crosslinker with an established role in cytokinesis is α-actinin (Fujiwara et al., 1978; Mukhina et al., 2007). Finally, actin filaments must be disassembled for successful completion of cytokinesis and crucial to this process is the actin severing protein ADF/cofilin, Twinstar (Tsr) in Drosophila (Gunsalus et al., 1995; Ono et al., 2003).

Non-muscle myosin II exists as a pair of heavy chains each associated with an essential light chain and a regulatory light chain. Upon activation, myosin II forms into filaments and uses its inherent ATPase activity to translocate antiparallel actin filaments, thereby generating force. Activation occurs by phosphorylation of myosin regulatory light chain, also known as Spaghetti squash (Sqh) in Drosophila (Yamakita et al., 1994; Jordan and Karess, 1997; Komatsu et al., 2000). There are three classes of myosin regulatory light chain kinases in animal cells – myosin light chain kinase (MLCK), Rho-dependent kinase (ROCK) and citron kinase, and there is some evidence that each contributes to cytokinesis (Matsumura, 2005). However, the evidence for MLCK is not as conclusive as for the other two. Like the kinases, myosin phosphatase (MYPT) must also be tightly regulated during cytokinesis. Although active earlier in mitosis, it becomes inactivated at the cleavage furrow during cytokinesis by phosphorylation of inhibitory sites (Kawano et al., 1999; Yokoyama et al., 2005). This inhibition may need to be relieved at the end of cytokinesis when the CR is disassembled.

1.1.2.2 Septins

Septins are a conserved family of GTP-binding proteins first discovered in the budding yeast, Saccharomyces cerevisiae, in a genetic screen for mutants defective in cytokinesis and cell morphology (Hartwell, 1971). Since then, septins have been identified and characterized in a wide assortment of animals and fungi (Weirich et al., 2008). The number of septin genes per organism varies greatly, from just two in C. elegans to fourteen in humans. In S. cerevisiae, septins localize to the mother-bud neck, where they form a filamentous collar and function in bud-site selection, spindle positioning, polarized growth and cytokinesis. In general, septins associate with both the plasma membrane and the actin and microtubule cytoskeletons. In addition to their conserved function in cell division, septins have been shown to have other functions in the nervous system and in the mammalian sperm tail annulus (Beites et al., 1999; Finger et al., 2003; Steels et al., 2007).

At the individual protein level, septins contain several conserved regions: an amino- (N-) terminal polybasic region required for phospholipid interactions (see Section 1.1.3); a GTP- binding domain belonging to the P-loop family of GTPases; and a variable coiled-coil region at the carboxy- (C-) terminus. Septins interact with each other through their coiled-coil regions to form hetero-oligomeric complexes containing two of each septin type (John et al., 2007; Sirajuddin et al., 2007; Bertin et al., 2008). The number of septins in the core complex can vary. In C. elegans, the complex contains two septins, whereas the core complex in Drosophila and humans generally contains three septins and fungal complexes can have four or five (Barral and Kinoshita, 2008). Oligomers can assemble into non-polar filaments and, at higher levels of organization, filaments form rings or gauzes (Kinoshita, 2003; Rodal et al., 2005).

In S. cerevisiae, septins are essential for cytokinesis (Hartwell, 1971). During budding, septins form a collar at the bud site that, at the onset of cytokinesis, splits into two separate rings (Lippincott et al., 2001; Dobbelaere and Barral, 2004). This rearrangement involves a dramatic 90˚ rotation of filaments, during which they shift from an hourglass-like formation to circumferential rings (Vrabioiu and Mitchison, 2006). Though highly dynamic during this period, septin filaments are immobile both before and after (Dobbelaere et al., 2003). A primary function for septins in this system is compartmentalization of the cortex at the site separating mother and bud. When the ring splits into two, septins form diffusion barriers on either side that

10 restrict many proteins to the neck area including: myosin II, the pombe Cdc15 homology (PCH) protein Hof1p, the IQ motif containing GTPase activating protein (IQGAP) homologue Ipq1p and members of the polarisome and exocyst complexes (Epp and Chant, 1997; Lippincott and Li, 1998; Korinek et al., 2000; Vallen et al., 2000; Iwase et al., 2007). The mechanism by which septins form this diffusion barrier is still not fully understood.

In contrast to S. cerevisiae, septins in S. pombe are not essential. The septin mutant, spn4Δ, displays a mild cytokinesis defect similar to that of the anillin-like protein Mid2p (see Section 1.2.3) (Longtine et al., 1996; Berlin et al., 2003; Tasto et al., 2003). Analysis of the spn4Δ phenotype suggests that S. pombe septins function in exocytosis and septum digestion.

Septins are essential to animal cell cytokinesis in many organisms. In C. elegans, Drosophila and mammalian cells, septins localize to the cleavage furrow (Fares et al., 1995; Kinoshita et al., 1997; Nguyen et al., 2000). Antibody injection or RNAi targeted against mammalian SEPT2 and SEPT9 leads to formation of binucleate cells (Kinoshita et al., 1997; Surka et al., 2002). In Drosophila, a null mutation in the septin gene peanut (pnut) results in the formation of imaginal tissues containing clusters of multinucleate cells (Neufeld and Rubin, 1994). In addition, when maternal contribution of peanut is eliminated, embryos exhibit defects in cellularization – a specialized form of cytokinesis (Adam et al., 2000). Unlike in Drosophila and mammalian cells, septins do not appear to be essential for cytokinesis in C. elegans; embryonic cells divide successfully when either or both of the worm septins is inhibited by RNAi or mutation (Nguyen et al., 2000; Maddox et al., 2005). Although not essential, septins act with anillin to promote asymmetric furrow ingression (see Section 1.2.3) (Maddox et al., 2007).

The role of septins during cytokinesis is not as clear in animal cells as it is in budding yeast. One likely function of septins is to act as a molecular scaffold within the cleavage furrow. The mammalian septin SEPT2 binds myosin II, and this association is required for myosin activation during cell division (Joo et al., 2007). Septins also interact indirectly with F-actin, using anillin as an adaptor (see Section 1.2.3) (Kinoshita et al., 2002). Additionally, septins may play a mechanical role in deforming the furrow membrane. In a recent study, the addition of septins to giant liposomes resulted in spontaneous tubulation in vitro, and this process was facilitated by the presence of phosphatidylinositol 4-phosphate (PI4P) or phosphatidylinositol

4,5-bisphosphate (PIP2) (Tanaka-Takiguchi et al., 2009). Finally, there is evidence that septins regulate membrane trafficking during late stages of cytokinesis. Mammalian SEPT5 binds to the t-SNARE syntaxin-1 and this interaction leads to inhibition of secretion (Beites et al., 1999; Dent et al., 2002). In a proposed model of septin function during late cytokinesis, septins delineate a permissive zone at the midbody where vesicle fusion contributes to successful abscission (Joo et al., 2005).

1.1.2.3 Membrane lipids

It is now well accepted that the lipid composition of the plasma membrane in the vicinity of the cleavage furrow is itself important for cytokinesis. For example, the integral membrane lipid phosphatidylethanolamine (PE) normally resides in the inner leaflet of the plasma membrane bilayer. During cytokinesis, PE is transferred to the external leaflet and this redistribution is required for disassembly of the CR and, thus, completion of cytokinesis (Emoto and Umeda, 2001). Very-long-chain fatty acids or their derivative complex lipids may be required for efficient cleavage during division. Mutations in the Drosophila gene bond – encoding a member of Elovl proteins that mediate elongation of very-long-chain fatty acids – severely slow furrow ingression in spermatocytes (Szafer-Glusman et al., 2008). Similarly, mutations in members of Drosophila peroxin (pex) family of genes, required for peroxisome biogenesis and very-long- chain fatty acid metabolism, also cause failures in male meiotic cytokinesis (Chen et al., 2010). Another recent study suggests that membrane subdomains, sometimes referred to as lipid rafts, play an important signalling role in dividing sea urchin eggs (Ng et al., 2005). These domains are enriched in sphingolipids, gangliosides and cholesterol, as well as tyrosine phosphorylated proteins. Likewise, sterol-rich membrane domains are localized to the site of division in S. pombe (Takeda et al., 2004) and, in Drosophila, a predicted sphingolipid delta-4 desaturase, Des-1, is required for central spindle assembly and cytokinesis (Basu and Li, 1998).

The most extensively studied class of membrane lipid with a clear contribution to cytokinesis is the phosphatidylinositol (PI) phosphates, or PIPs (reviewed in Janetopoulos and

Devreotes, 2006; Logan and Mandato, 2006). In Drosophila and mammalian cells, PIP2 is found in the cleavage furrow and is required for cytokinesis (Field et al., 2005b; Wong et al., 2005). In

Dictyostelium, local production of phosphatidylinositol 3,4,5-trisphosphate (PIP3) at the poles of the cell also contributes to successful cleavage (Janetopoulos et al., 2005). Multiple PIP2-

12 binding proteins are hypothesized to be involved in making connections between the furrow membrane and the underlying cytoskeleton (discussed further in Section 1.1.3). Sequestration of

PIP2 using specific lipid-binding domains interferes with the connection between the CR and the furrow membrane (Field et al., 2005b; Wong et al., 2005). In addition, drugs that interfere with

PIP2 hydrolysis cause cleavage furrow instability and eventual regression, suggesting that PIP2 must be removed for successful completion of cytokinesis (Saul et al., 2004; Wong et al., 2005;

Naito et al., 2006). Alternatively, or in addition, PIP2-derived second messengers could be required for cytokinesis.

Enzymes that affect the distribution or phosphorylation state of phosphatidylinositol have also been implicated in cytokinesis. A phosphatidylinositol transfer protein, presumably responsible for getting PI lipids to the equatorial membrane, is essential for cytokinesis in Drosophila (Gatt and Glover, 2006; Giansanti et al., 2006). Kinases that act on PI and its derivatives have been shown to play direct roles in cytokinesis in S. pombe, Dictyostelium and Drosophila (Brill et al., 2000; Zhang et al., 2000; Janetopoulos et al., 2005). In S. pombe, a phosphatidylinositol-4-phosphate 5-kinase (PIP5K) homologue, Its3, colocalizes with PIP2 at the furrow (Zhang et al., 2000). its3 temperature sensitive mutants exhibit an increased septation index at the restrictive temperature, indicative of cytokinesis failures. In Dictyostelium, phosphatidylinositol-3 (PI3) kinases are required for PIP3 generation at the poles (Janetopoulos et al., 2005). Disruption of PI3 kinases, either by mutation or chemical inhibitors, results in formation of multinucleate cells. Finally, in Drosophila, the phosphatidylinositol 4-kinase (PI4K) IIIβ, also known as Four-wheel drive (Fwd), is essential for male meiotic cytokinesis (Brill et al., 2000). Redundancy with other PI4Ks may explain why Fwd is not required for cytokinesis in all cells. Alternatively, the importance and relative contribution of PIPs to successful cytokinesis may vary across cell types. Although Fwd could generate PI4P as a substrate for PIP2 synthesis in the cleavage furrow, its localization to the Golgi and its interaction with the trafficking protein Rab11 suggest that Fwd and PI4P contribute to cytokinesis by affecting intracellular membrane trafficking pathways (e.g., secretion, recycling) (Polevoy et al., 2009). The PI lipid phosphatidylinositol-3-phosphate (PI3P), usually associated with endosomal compartments, also contributes to cytokinesis in mammalian cells by recruiting CHMP4B, a member of the ESCRT-III complex, to the midbody (Sagona et al., 2010). Thus, the role for PIPs in cytokinesis is not spatially restricted to the plasma membrane.

1.1.2.4 Rho GTPase, regulators and effectors

Members of the Rho family of GTPases, which includes Rho, Rac and Cdc42, are important regulators of the actin cytoskeleton in a variety of cell types and processes (reviewed in Jaffe and Hall, 2005). It is now well established that the Rho GTPase, RhoA, is a master regulator of animal cell cytokinesis (Piekny et al., 2005). Active RhoA localizes to the cleavage furrow in dividing echinoderm and amphibian embryos (Bement et al., 2005; Yuce et al., 2005) and its downstream effectors are responsible for multiple aspects of CR function, including F-actin polymerization and myosin activation.

RhoA can exist in either an active GTP-bound form or an inactive GDP-bound form. Specific regulatory enzymes affect the state of RhoA – GDP-GTP exchange factors (GEFs) promote GTP binding and GTPase activating proteins (GAPs) promote GTP hydrolysis. During cell division, the RhoGEF (ECT2 in mammals, Pebble (Pbl) in Drosophila) is responsible for activating RhoA. Disruption of ECT2/Pbl leads to a failure in cytokinesis (Prokopenko et al., 1999; Tatsumoto et al., 1999). Proper localization of ECT2/Pbl depends on a complex termed centralspindlin, composed of the kinesin MKLP1 (Pavarotti (Pav) in Drosophila) and the Rho- family GAP MgcRacGAP (RacGAP50C in Drosophila) (Somers and Saint, 2003; Yuce et al., 2005). It remains unresolved whether the GAP activity of MgcRacGAP acts on RhoA and/or Rac during cytokinesis (Canman et al., 2008; Glotzer, 2009; Miller and Bement, 2009). Centralspindlin is thought to travel along spindle microtubules using the motor activity of MKLP1/Pav. The complex becomes highly localized at the presumptive site of division where MgcRacGAP/RacGAP50C (hereafter referred to as RacGAP) physically interacts with ECT2/Pbl, leading to RhoA activation.

The effectors of RhoA with established roles in cytokinesis include the formin, mDia2/Dia, and the myosin light chain kinases, ROCK and citron kinase. As discussed in Section 1.1.2.1, these proteins are responsible for F-actin polymerization and myosin activation, respectively.

1.1.2.5 Kinases, ubiquitin ligases, trafficking proteins and more

Aside from the core molecules already discussed, many other proteins have well-established roles in cytokinesis. These include kinases, trafficking proteins and ubiquitin ligases. In brief,

14 the cell cycle regulator Cdk1/cyclin B1 must be inactivated during anaphase for cytokinesis to initiate. Cdk1 has been shown to phosphorylate and negatively regulate several proteins involved in cytokinesis, including MKLP1 and ECT2 (Mishima et al., 2004; Yuce et al., 2005). In contrast, Aurora B kinase and PLK1 (Polo in Drosophila) have activating roles in cytokinesis. Aurora B is part of a complex termed the chromosomal passenger complex, which, during cell division, travels from chromosomes to centromeres and then to the central spindle. Among the substrates for Aurora B are MKLP1 and RacGAP, the two components of centralspindlin (Minoshima et al., 2003; Guse et al., 2005). Like Aurora B, PLK1 associates with chromosomes and the central spindle and, in addition, localizes to centrosomes. Phosphorylation of MgcRacGAP by PLK1 is required for recruitment of ECT2 to the central spindle (Burkard et al., 2007; Petronczki et al., 2007; Burkard et al., 2009; Wolfe et al., 2009). The net result is that both Aurora B and Polo-like kinase feed into the RhoA activation pathway, leading to actomyosin contractility and furrow ingression.

As previously described, membrane trafficking is important during cytokinesis, especially during very late stages. Accordingly, many proteins involved in this process have been identified in RNAi or genetic screens. Fwd and Rab11 are two examples already discussed. Additional trafficking proteins with roles in cytokinesis include members of the exocyst and ESCRT complexes, SNARE membrane fusion proteins and other Rab family GTPases (reviewed in Barr and Gruneberg, 2007; Montagnac et al., 2008).

Finally, protein degradation is required for successful completion of cytokinesis and the APC/C ubiquitin ligase is involved in this process. Some of the many targets of APC/C include PLK1 and anillin (see below) (Lindon and Pines, 2004; Zhao and Fang, 2005).

The proteins and lipids discussed above have well-established, conserved functions in animal cell cytokinesis. However, this is by no means a complete list. Cytokinesis is a complex process with redundant or overlapping mechanisms in place to ensure robustness. Thus, there are likely to be additional proteins that, although contributing to the process, would not be identified in single mutant or knockdown experiments. These might include additional trafficking proteins or proteins that interact with F-actin, microtubules and/or the plasma membrane.

1.1.3 Membrane-cytoskeletal attachments in cytokinesis

One feature of animal cell cytokinesis that remains poorly understood is the nature of the attachment of the CR to the overlying plasma membrane. The relationship between the CR and the membrane is inherently complex; on the one hand, the attachment must be strong enough to deform the membrane initially and to counteract membrane tension as the furrow ingresses. On the other hand, the CR must be disassembled as it shrinks, and thus the attachment cannot be too strong. Furthermore, components of the contractile ring may be quite dynamic (as discussed in Section 1.1.1.2), with filaments assembling as well as disassembling. If this is the case, the connection between the CR and the membrane must be even more flexible. How the cell maintains this delicate balance between too strong an attachment and one that is not strong enough is still largely unknown.

There are a number of candidate proteins that could in theory provide a link between the CR and the furrow membrane. Within the membrane, one of the major targets of these interactions appears to be the lipid PIP2. Indeed, there is an intimate relationship between plasma membrane PIP2 and the actin cytoskeleton (Logan and Mandato, 2006; Saarikangas et al., 2010). Proteins such as those of the ezrin/radixin/moesin (ERM) family (discussed below) provide structural links between PIP2 and F-actin. Additionally, PIP2 is involved in regulating F- actin through activation of the Arp2/3 complex via N-WASp and inhibition of cofilin and the capping protein, gelsolin (Yin and Janmey, 2003).

Two potential candidates for linking the actomyosin ring and the membrane during cytokinesis are septins and anillin. The polybasic region of septins can interact with phosphoinositides in mammalian cells and budding yeast. For example, the mammalian septin,

SEPT4, binds preferentially to PIP2 and PIP3 in vitro and treating cells with agents that occlude, dephosphorylate or hydrolyze PIP2 alters SEPT4 localization (Zhang et al., 1999). In addition, a recombinant septin complex of SEPT2/6/7 has the ability to associate with and deform liposomes containing PI4P or PIP2 (Tanaka-Takiguchi et al., 2009). The S. cerevisiae septin Cdc11 shows preferential binding to PI4P and phosphatidylinositol 5-phosphate (PI5P) in vitro, and cells with temperature-sensitive alleles in either of the two essential yeast PI4Ks, stt4 or pik1, show delocalization of a Cdc11-GFP construct (Casamayor and Snyder, 2003). Furthermore, depletion of PIP2 by overexpression of yeast PI3K causes dramatic rearrangement of septin filaments

(Rodriguez-Escudero et al., 2005). Anillin, which links septins to F-actin (Kinoshita et al., 2002), may also interact with PIPs through its conserved pleckstrin homology (PH) domain (see Section 1.2.1).

A number of other membrane-cytoskeleton crosslinkers are candidates to regulate cytokinesis. ERM proteins, which consist of an N-terminal phospholipid-binding FERM domain and a C-terminal F-actin-interacting region, localize to the cleavage furrow in Drosophila and in mammalian cells (Sato et al., 1991; Bretscher et al., 2002; Carreno et al., 2008; Kunda et al., 2008). In S. cerevisiae, the C2 lipid-binding domain-containing protein Inn1 colocalizes with the actomyosin ring and is required for cytokinesis (Sanchez-Diaz et al., 2008). It remains to be seen whether any C2-domain-containing proteins function similarly in metazoan cells. Other proteins could contribute to CR-membrane attachment by interacting with phospholipids other than PIPs or by binding transmembrane proteins. The actin-binding protein, talin, which interacts with integrins, localizes to the midbody in mammalian cells and could be an example of the latter (Bellissent-Waydelich et al., 1999). It seems likely that CR-membrane attachment during cytokinesis requires the coordination of multiple interactions involving a number of proteins, lipids and complexes.

Outside of the context of cytokinesis, a set of proteins with a well-established role in linking the plasma membrane and the actin cytoskeleton are components of adherens junctions (Pokutta and Weis, 2007; Hartsock and Nelson, 2008). Classical cadherins, such as E-cadherin, are transmembrane proteins. Cadherins in adjacent cellular membranes are able to form Ca2+- mediated, homophilic interactions. Within the cytoplasm, cadherins associate with proteins termed catenins. Based on early biochemical data, E-cadherin was originally thought to bind directly to β-catenin, which binds to α-catenin, which in turn binds to F-actin (Aberle et al., 1994; Rimm et al., 1995). The picture became more complex with the finding that α-catenin can bind to either β-catenin or F-actin, but not both simultaneously (Yamada et al., 2005). This result suggests that the association between adherens junctions and F-actin is more dynamic than once thought. Currently, there are no reports of direct contributions by cadherin-catenin complexes to the process of cytokinesis. However, results presented in Chapter 3 strongly suggest such a role may exist. An in-depth discussion of possible connections between adherens junction proteins and cytokinesis, and the implications thereof, will be presented in Chapter 4.

1.1.4 The current cytokinesis picture: complexity, redundancy and open questions

Recent reviews suggest a general molecular model of cytokinesis that can apply to almost all dividing animal cells (Glotzer, 2005; Eggert et al., 2006; Barr and Gruneberg, 2007). The process is initiated by key regulatory kinases: Cdk1 is degraded, whereas Aurora B and PLK1 are activated. The centralspindlin complex, upon activation, travels along a subset of microtubules to the presumptive site of division, where interactions with ECT2/Pbl lead to RhoA activation. RhoA acts through multiple effectors to assemble and initiate constriction of a contractile ring containing F-actin and myosin II. Anillin, septins and additional actin-binding proteins associate with the CR, providing structural and regulatory support. The CR constricts and robust interactions with PIPs and other membrane components result in cleavage furrow ingression. The furrow ingresses until an intercellular bridge and midbody are formed. The CR is disassembled and ubiquitination of several proteins by APC/C marks them for degradation. Membrane trafficking and membrane fission contribute to final separation of daughter cells by abscission.

The available data go a long way toward answering many of the central, outstanding questions in the field, but the model is far from complete. Focusing on any part of the process, the complexity of the system quickly becomes evident. Different combinations of molecular interactions and built-in redundancies have evolved to ensure that cells can divide successfully under a range of conditions. Improved future models of cytokinesis will need to take into account this complexity and subtlety. Many questions remain to be answered, including how dynamic or stable components of the CR are and how the CR maintains its attachment to the membrane. Within this developing model of cytokinesis, some proteins appear to be crucial in holding the various proteins and processes together. The evolutionarily conserved protein anillin is emerging as one of the key factors responsible for this coordination.

1.2 Anillin and anillin-related proteins

Anillin is a conserved, multidomain scaffolding protein first discovered in Drosophila through F- actin affinity chromatography experiments performed on embryonic extracts (Miller et al., 1989). It was given the name anillin, derived from the Spanish word for ring, due to its contractile ring localization in dividing cells (Field and Alberts, 1995). anillin alleles were

18 initially identified as alleles of the gene scraps (scra) in a screen for recessive female-sterile mutations (Schupbach and Wieschaus, 1989). Anillin was later confirmed as the product of scra (Field et al., 2005a). Anillin homologues have been identified in numerous organisms from yeast to humans (reviewed in D'Avino, 2009). Experiments employing genetics, RNAi techniques or antibody injections clearly demonstrate an essential role for anillin in cytokinesis (Oegema et al., 2000; Somma et al., 2002; Kiger et al., 2003; Echard et al., 2004; Eggert et al., 2004; Field et al., 2005a). Although progress has been made in our understanding of anillin function, its precise contribution during different stages of cytokinesis remains poorly understood. In addition, the finding that anillin is highly expressed in the mammalian central nervous system (CNS) hints at potential roles for anillin outside of cytokinesis (Hall et al., 2005b). Indeed, recent reports suggest a link between anillin and cancer (Hall et al., 2005b; Suzuki et al., 2005; Olakowski et al., 2009). Hence, an improved understanding of anillin function may thus lead to the development of novel anti-cancer therapies.

In this section I will summarize what is currently known about anillin and anillin-related proteins from studies in diverse organisms. I will begin with an overview of anillin protein structure, highlighting conserved and non-conserved regions across species.

1.2.1 Anillin structure

Anillin or anillin-related proteins have been identified in many animals and fungi. The number of homologues per organism varies: Drosophila and vertebrates have a single anillin, whereas C. elegans has three. There are two well-characterized anillin-related proteins in S. pombe, which have a number of similarities to metazoan anillins. S. cerevisiae also appears to have two anillin- related proteins, although the functional similarities in this organism are less clear. The only conserved region across all anillin-like proteins is a PH domain at the C-terminus. Several other structural regions are conserved across a subset of anillin homologues.

The Drosophila anillin contains 1239 amino acids and has an apparent mobility of approximately 190 kilodaltons (kDa) on SDS-polyacrylamide gel electrophoresis (SDS-PAGE) (Field and Alberts, 1995). Like other anillin proteins, Drosophila anillin contains a PH domain near the C-terminus (Figure 1.2). A region just upstream of the PH domain with high homology among metazoan anillins is referred to as the anillin homology region (AHR) (Oegema et al., 2000). Closer to the N-terminus is an actin-binding region, which, with short flanking

19 sequences, is also responsible for bundling actin filaments (Field and Alberts, 1995). Several Src-homology 3 (SH3) binding motifs and nuclear localization signals (NLS) are predicted from the amino acid sequence. An NLS located just upstream of the PH domain was shown to bind directly to nuclear importins (Silverman-Gavrila et al., 2008). In addition, a region of anillin upstream of the PH domain and likely within the AHR is responsible for an interaction with RacGAP50C (D'Avino et al., 2008; Gregory et al., 2008).

Figure 1.2. Schematic representation of the Drosophila anillin protein.

Anillin is a multidomain scaffolding protein containing 1239 amino acids (aa). Indicated domains or motifs are required for protein-protein interactions and/or anillin localization. Asterisks indicate amino acid sequences corresponding to interacting regions identified for vertebrate anillin. These have yet to be confirmed in Drosophila. Arrows point to short stretches of amino acids required for APC/C or RhoA binding.

Vertebrate anillin is similar to Drosophila anillin in both structure and function. It also contains a PH domain and AHR at its C-terminus, an actin-binding region toward the N-terminus and predicted SH3-binding sites and NLSs. In contrast to Drosophila anillin, an N-terminal region of human anillin containing three putative NLSs is responsible for nuclear localization (Oegema et al., 2000). Xenopus laevis anillin was shown to bind phosphorylated myosin II through a region located just upstream of the actin binding region (Straight et al., 2005). This

20 myosin-binding region contains several amino acids that are invariant across human, Xenopus and Drosophila anillins. Consistent with this, myosin was identified as a binding partner of Drosophila anillin by mass spectrometry (D'Avino et al., 2008). It is yet to be established whether human anillin binds RacGAP; however, an additional interaction was recently reported between human anillin and RhoA via a short sequence in the AHR (Piekny and Glotzer, 2008). Mammalian anillin also contains a destruction-box sequence near the N-terminus that is required for ubiquitination by APC/C (Zhao and Fang, 2005). Ubiquitination results in degradation of anillin at the end of cell division. It is not known whether Drosophila anillin is also ubiquitinated and degraded, although similar profiles of protein levels over the cell cycle suggest that this is likely the case (Field and Alberts, 1995; Zhao and Fang, 2005).

The three C. elegans anillin homologues, referred to as ANI-1, ANI-2 and ANI-3, all contain a C-terminal PH domain and an AHR (Maddox et al., 2005). ANI-1 is the closest in overall conservation to Drosophila and human anillin in that it contains regions with sequence similarities to the actin and myosin binding regions. No conserved domains or motifs have been described within the N-terminal regions of ANI-2 or ANI-3.

The evolutionary relationship between yeast anillin-like proteins and metazoan anillins is not entirely clear. Two strong candidates for anillin-related proteins in yeast are Mid1p and Mid2p in S. pombe. Like metazoan anillins, Mid1p and Mid2p have PH domains at their C- termini. Mid1p contains an NLS site as well as two nuclear exit signals (NES), both of which are required for proper localization (Paoletti and Chang, 2000). Like Xenopus anillin, Mid1p interacts with fission yeast myosin-II heavy chain, Myo2 (Motegi et al., 2004). However, the region of Mid1p required for this interaction has not been mapped. Additionally, Mid1p also has a defined region required for interacting with Clp1/Flp1, a Cdc14 phosphatase homologue that regulates CR component dynamics (Clifford et al., 2008). In addition to the PH domain of Mid2p, the only other characterized Mid2p sequence is an N-terminal region that contains three putative PEST (Pro/Glu/Ser/Thr) domains required for ubiquitination and degradation (Tasto et al., 2003).

It has been proposed that two proteins, Boi1 and Boi2, are S. cerevisiae anillins (Norden et al., 2006). Along with SH3 domains, Boi1 and Boi2 have C-terminal PH domains, and the PH domain of Boi1 was shown to interact with PIP2 (Hallett et al., 2002). The functional connection

21 between Boi1/Boi2 and other anillins is less clear; although Boi1 and Boi2 localize to the bud site in dividing cells, they were shown to play inhibitory rather than activating roles in cytokinesis (Norden et al., 2006). Because of this divergence, Boi1/Boi2 will not be discussed further here.

1.2.2 Anillin expression and localization

The single anillin in Drosophila and vertebrates is widely expressed. In Drosophila, anillin is expressed throughout development and in tissue culture cells (Field and Alberts, 1995). Similarly, in human tissue profiling experiments, anillin was detected in a wide variety of tissues (Hall et al., 2005b). Drosophila tissue culture cells that have exited the cell cycle (i.e., that are in

G0) have no detectable anillin, suggesting that anillin expression may be limited to dividing cells (Field and Alberts, 1995). This is contradicted, however, by the presence of high levels of anillin in the human CNS – a region with very low levels of cell division (Hall et al., 2005b).

In dividing cells, Drosophila and vertebrate anillin have the same localization pattern (Field and Alberts, 1995; Oegema et al., 2000; Straight et al., 2005): during interphase, anillin localizes to the nucleus; during metaphase, it relocalizes to the cell cortex; and shortly thereafter becomes enriched at the equator. Anillin remains in the cleavage furrow, associated with the CR, until completion of cytokinesis. At the end of cell division, anillin is degraded and new protein is synthesized during the next cell cycle (Field and Alberts, 1995; Zhao and Fang, 2005).

In addition to its localization to the cleavage furrow in cells undergoing conventional cytokinesis, Drosophila anillin localizes to other related structures. It is found in metaphase furrows and at the cellularization front in embryos undergoing cellularization (Field and Alberts, 1995), a process with strong similarities to conventional cytokinesis (Mazumdar and Mazumdar, 2002). In addition, anillin localizes to stable intercellular bridges called ring canals that form during incomplete cytokinesis in developing germ cells (see Section 1.3.1) (Hime et al., 1996; de Cuevas and Spradling, 1998; Giansanti et al., 1999).

Anillin localization in C. elegans suggests specialized functions for the different homologues. ANI-1 and ANI-2 are expressed in largely non-overlapping tissues and developmental stages (Maddox et al., 2005). ANI-1 is predominantly expressed in the embryo, where it concentrates in cortical patches during contractile events known as membrane ruffling,

22 pseudo-cleavage and polar body extrusion. It also localizes to cleavage furrows during early embryonic cytokinesis. In contrast, ANI-2 is predominantly expressed in the adult worm. Within the hermaphrodite gonad, ANI-2 localizes to the lining of a structure called the rachis, which contains the cytoplasmic core of the gonad. The subcellular localization of ANI-3 has not been characterized.

The localization of Mid1p in S. pombe is similar, but not identical, to fly and vertebrate anillin. In interphase cells, Mid1p primarily localizes to the nucleus, with a subset of the protein found in spots over a broad region in the medial cell cortex (Sohrmann et al., 1996; Bahler et al., 1998; Paoletti and Chang, 2000). After mitotic entry, Mid1p is exported from the nucleus and becomes the first core cytokinesis-related protein detected at the equator (Wu et al., 2003). It remains associated with the equator until septum formation, at which point it is transported back into the nucleus. Mid2p joins the medial ring much later than Mid1p and associates with septins (Berlin et al., 2003; Tasto et al., 2003). After division is complete, Mid2p is ubiquitinated and degraded (Tasto et al., 2003).

1.2.3 Known and predicted functions of anillin

The function of Drosophila and vertebrate anillin during cytokinesis appears to be highly conserved. Antibody injections or RNAi knockdowns performed in various tissue culture cell lines leads to the formation of multinucleate cells, revealing that anillin is essential for this process (Oegema et al., 2000; Somma et al., 2002; Echard et al., 2004; Straight et al., 2005; Zhao and Fang, 2005; D'Avino et al., 2008; Piekny and Glotzer, 2008). The effect of anillin knockdown only becomes visible during late stages of cytokinesis. That is, a cleavage furrow forms normally and ingresses, but eventually becomes unstable and regresses. This runs counter to predictions that anillin would be required for formation of the CR, based on its early localization to the furrow and its ability to bind and bundle F-actin (Field and Alberts, 1995; Giansanti et al., 1999). Instead, loss of anillin is associated with delocalized actomyosin contractility, with myosin regulatory light chain moving around the periphery of the cell rather than remaining concentrated at the equator (Straight et al., 2005; Hickson and O'Farrell, 2008; Piekny and Glotzer, 2008).

Anillin has the ability to localize septins. In mammalian cells, expression of a C-terminal portion of human anillin, which includes the PH domain plus a small amount of upstream

23 sequence, leads to formation of ectopic cortical foci containing the anillin construct and the septin SEPT7 (Oegema et al., 2000). In in vitro assays, anillin is required as an adaptor to link septins and F-actin (Kinoshita et al., 2002). In Drosophila, embryos derived from mothers homozygous for anillin alleles with mutations in the PH domain show defects in peanut localization to the cellularization front (Field et al., 2005a). Moreover, a C-terminal anillin fragment was shown to copurify with peanut and Sep2 (D'Avino et al., 2008) and mass spectrometry of anillin-interacting proteins in Drosophila identified three septins – peanut, Sep2 and Sep5 (D'Avino et al., 2008; Silverman-Gavrila et al., 2008).

Anillin plays an additional role in linking important signalling molecules to the CR. In Drosophila cells, interdependent localization of RacGAP50C and anillin at the cleavage furrow has been proposed to serve as a crucial intermolecular link between spindle microtubules and the contractile ring (D'Avino et al., 2008; Gregory et al., 2008). Anillin also interacts with RhoA and is required for proper RhoA localization at the furrow (Zhao and Fang, 2005; Piekny and Glotzer, 2008). In addition, anillin localization is at least partially dependent on Pbl, likely acting through RhoA (Prokopenko et al., 1999; Hickson and O'Farrell, 2008).

Since cytokinesis is an essential developmental process and anillin is required for cytokinesis, other roles for anillin may be obscured. However, there is already evidence that anillin functions in processes other than conventional cytokinesis. In Drosophila, anillin is required for cellularization as well as somatic cytokinesis (Field et al., 2005a). A class of alleles with mutations in the PH domain causes defects in the timing and rate of furrow ingression during cellularization. Moreover, membrane stability appears to be affected, as dramatic vesiculation is seen between newly formed membranes. In addition, a separate mutation closer to the N-terminus causes defects in a specialized type of cytokinesis called pole cell formation, in which progenitors of germ cells pinch off the posterior end of the syncytial embryo. In a separate study of a zygotic anillin allele, anillin was shown to be required for asymmetric cell divisions of the larval peripheral nervous system (PNS), although it is not clear whether anillin plays a functional role in setting up the asymmetry (O'Farrell and Kylsten, 2008). Finally, anillin is highly expressed in the mammalian CNS (Hall et al., 2005b). Because of its ability to bind key cytoskeletal components and regulators, it is possible that anillin may have evolved additional functions, for example, acting as a scaffold in the formation of actin rich structures within neurons.

The C. elegans anillin homologues appear to carry out specialized, non-overlapping functions. ANI-1 is required for contractile events in the early embryo, including cortical ruffling, pseudocleavage and polar extrusion (Maddox et al., 2005). In early embryonic cytokinesis, ANI-1, in concert with the septins, promotes a symmetry-breaking mechanism that is proposed to make cytokinesis more robust in metazoan cells (Maddox et al., 2007). Like Drosophila and vertebrate anillin, ANI-1 targets the septins, but not myosin II, to the contractile ring. Surprisingly, ANI-1 is dispensable for cytokinesis later in embryonic development. ANI- 2, like ANI-1, contributes to embryonic viability, although its function appears to be limited to the rachis in the hermaphrodite gonad (Maddox et al., 2005). ANI-2 is proposed to play a structural role, in that its depletion leads to premature disconnection of oocytes and generation of embryos of varying sizes. In contrast to its homologues, ANI-3 is dispensable for embryonic viability and has no established function.

The two anillin-like proteins in S. pombe carry out functions performed by the single anillin in Drosophila and vertebrates, although there are notable differences. Some of these are likely connected to differences in the process of cytokinesis itself. For example, in S. pombe, Mid1p plays a central role in connecting the nucleus with the overlying cortex and establishing the site of division. The polo-like kinase, Plo1p, phosphorylates Mid1p and is responsible for its exit from the nucleus (Bahler et al., 1998). The cell-polarity kinase Pom1p keeps Mid1p focused at the midzone by negatively regulating it at the poles (Celton-Morizur et al., 2006; Padte et al., 2006). Unlike human anillin, the PH domain of Mid1p is not required for localization to the cortex (Paoletti and Chang, 2000). Instead, cortical localization depends on two independent protein regions, one C-terminal and one N-terminal, each capable of targeting protein fragments to the cortex. An amphipathic helix located near the C-terminus is predicted to insert directly into the phospholipid bilayer, whereas an N-terminal fragment is thought to localize to the cortex via interactions with other cortical proteins (Paoletti and Chang, 2000; Celton-Morizur et al., 2004). Once at the medial cortex, Mid1p is required for proper localization of myosin II heavy chain and the Cdc14 phosphatase Clp1 (Motegi et al., 2004; Clifford et al., 2008). In a prevalent model of S. pombe contractile ring formation, Mid1p establishes cortical nodes that eventually coalesce into an active actomyosin ring that encircles the cell (Wu et al., 2006). The anillin-like protein, Mid2p, which has been less intensively studied than Mid1p, primarily functions in

25 stabilizing septin rings late in cell division. Similar to septin mutants, mid2Δ mutants exhibit normal contractile ring closure and septum formation, but are defective in cell-cell separation.

In conclusion, active investigation over the last fifteen years has established anillin and anillin-related proteins as central players in animal and fungal cell cytokinesis. Indeed, in Drosophila and vertebrates, anillin is essential for cytokinesis and can interact with many core components of the cytokinetic machinery. Despite these findings, the functional contribution of anillin to cytokinesis and its regulation in time and space remain poorly understood. In addition, and as discussed above, recent results suggest the possibility that anillin contributes to cellular processes other than conventional cytokinesis. Drosophila spermatogenesis provides an excellent system in which to probe these aspects of anillin function.

1.3 The system: Drosophila spermatogenesis

Sperm development, or spermatogenesis, in Drosophila is a well-characterized system that encompasses many conserved elements, from stem cell maintenance to cell motility (Fuller, 1993, 1998). In this section, I provide a brief overview of the process, followed by more detailed information on aspects later referred to within this thesis – namely meiotic cytokinesis and individualization of mature sperm.

1.3.1 Overview

Spermatogenesis takes place within the Drosophila testis and initiates during larval development (Figure 1.3A). The process begins at the apical end of the testis at an epithelial structure called the hub, which is associated with germline and somatic stem cells. Each germline stem cell divides asymmetrically to give rise to a gonial cell and another stem cell. Concomitant with this, two cyst cells, themselves derived from the asymmetric division of somatic stem cells, envelop each gonial cell and continue to surround the gonial cell daughters as they divide and differentiate. The majority of development occurs within this unit, which is referred to as a cyst.

Each gonial cell undergoes four consecutive rounds of mitosis, giving rise to a group of sixteen primary spermatocytes. During the mitotic and subsequent meiotic divisions, the cells undergo a specialized, incomplete form of cytokinesis. After the actomyosin ring constricts to a diameter of 1-1.5 µm, the opening between daughter cells is stabilized and an intercellular bridge, termed a ring canal, is formed (Hime et al., 1996). In contrast to mature ring canals in

26 the female reproductive system, male germline ring canals are smaller, lack F-actin and contain anillin and septins (Hime et al., 1996; Giansanti et al., 1999). Male germ cells remain connected by ring canals throughout most of spermatogenesis, and thus develop within a syncytium.

Each cyst of sixteen Drosophila primary spermatocytes undergoes an extended period of cell growth, in which the cells increase approximately 25-fold in volume. This is followed by two rounds of meiotic division (discussed in Section 1.3.2) to form a cyst of 64 haploid spermatids. The spermatids then differentiate in a process termed spermiogenesis. The cells become polarized with the haploid nuclei clustered at one end of the cyst and sperm tails elongating in the opposite direction. During elongation, the sperm tails grow to an astonishing length of nearly 2 millimetres. Elongation is followed by an actin-dependent process called individualization (discussed in Section 1.3.3), in which each cell is invested with its own membrane. Individualized, motile sperm are transferred from the testis to the seminal vesicle and eventually to the female fly, where fertilization occurs. The entire process of spermatogenesis takes place over a ten day period at 25°C. During the life of the fly, the process is continuously restarted and each testis contains cysts at different stages of development.

Figure 1.3. Overview of Drosophila spermatogenesis and spermatid individualization.

(A) Schematic representation of Drosophila spermatogenesis beginning with germline stem cells (‘S’) (top) and culminating in the formation of mature sperm (bottom). Germline stem cells surround the apical hub (‘H’) and divide asymmetrically to give rise to new stem cells and founder gonial cells (‘G’). Each gonial cell undergoes four mitotic amplification divisions to produce a cyst of sixteen primary spermatocytes, which undergo sequential meiotic divisions to generate 64 haploid spermatids. Note that, starting with “Growth and Gene Expression”, a single spermatocyte and its meiotic products are shown. In the bottom two images, under “Spermatid Differentiation”, groups of cells are depicted with nuclei on the left and tails to their right. During much of development, male germ cells remain connected by stable intercellular bridges called ring canals (green) that form during mitotic and meiotic cytokinesis. Hence, male germ cells develop within a syncytial cyst. (B) Schematic representation of individualization. Individualization, which occurs during spermatid differentiation, is the process whereby 64 elongated spermatids (three representative cells are depicted) transition from being connected within a cyst to becoming autonomous sperm. (Left) Actin-based investment cones (red) form around nuclei (blue) and translocate along microtubule-based axonemes (green). (Right) As investment cones progress, they remove waste – pushed forward in a cystic bulge – and invest each spermatid with its own membrane. The atypical myosin, myosin VI, localizes to the leading edge of investment cones (yellow), and is required for their movement. Diagram in (A) was kindly provided by Margaret T. Fuller and modified by Julie A. Brill. Diagram in (B) was modified from a diagram kindly provided by Tatsuhiko Noguchi and Kathryn G. Miller.

1.3.2 Meiotic cytokinesis

Drosophila male meiosis is well established as one of the favoured model systems for studying cytokinesis in the fly (Fuller, 1993; Giansanti et al., 2001). Male germ cells have a number of obvious morphological landmarks when viewed by phase-contrast microscopy, making the stages before, during and after division easy to identify (Figure 1.4). Spermatocytes in early stages of meiosis are characterized by the presence of phase-dark mitochondria, which line up along the meiotic spindle. A cleavage furrow appears at the equatorial cortex at the midzone and eventually bisects the cell. One round of meiotic division is followed closely by the next and the volume of the cell is halved each time. After meiosis is complete, early round spermatids are easily identified by the presence of two discs of approximately equal size: a phase-light nucleus and a phase-dark mitochondrial derivative, which is formed by the aggregation and fusion of the cell’s mitochondria. This morphological feature provides a read-out for cytokinesis failure. In spermatids where cytokinesis has failed, multiple nuclei accompany enlarged mitochondrial derivatives that result from aggregation and fusion of mitochondria that should have been apportioned to more than one spermatid. By examining spermatids from flies carrying specific genetic mutations, one can easily determine whether cytokinesis failures have occurred and at what frequency.

Figure 1.4. Drosophila male meiotic cytokinesis.

(A-D) Phase contrast micrographs of wild-type (A-C) and fwd (D) male germ cells. (A) Example of a primary spermatocyte with large phase-light nucleus. (B) During meiotic cytokinesis, spermatocytes are bisected by cleavage furrows (arrows). Note the appearance of phase-dark mitochondria which line the meiotic spindle. (C) Post-meiotic early round spermatids contain a phase-light nucleus (arrowhead) and phase-dark mitochondrial derivative (arrow) of similar size. (D) Multinucleate early round spermatid from a fwd mutant. The presence of four nuclei (example, arrowhead) and enlarged mitochondrial derivative (arrow) indicate cytokinesis failures in both rounds of meiotic division. Figure provided by Julie A. Brill.

A number of features contribute to make male meiotic cytokinesis a useful system for studying animal cell cytokinesis. First, as described above, dividing spermatocytes have easily discernable structures and multinucleate cells are easy to identify. Second, cytokinesis in Drosophila spermatocytes has many similarities to other models of animal cell cytokinesis, both mechanically and genetically. The cells divide symmetrically and there is an intimate connection between spindle microtubules and the contractile ring (Giansanti et al., 2004). Moreover, many genes encoding core components of the conserved cytokinetic apparatus are required for spermatocyte cytokinesis. One notable difference between Drosophila spermatocytes and other cytokinesis models is the formation of stable ring canals, which occurs in place of abscission. However, several genes required for late stages of cytokinesis in other cells, including genes required for membrane trafficking, are also required in spermatocytes. Third, powerful genetic and molecular tools available to the Drosophila researcher can be applied to this system. For example, genetic screens have identified numerous mutants defective in male meiotic cytokinesis (Giansanti et al., 2004; Wakimoto et al., 2004). In addition, researchers have access to large collections of fly stocks and can take advantage of techniques to express tagged proteins or generate clones of mutant cells. Fourth, spermatocyte cytokinesis is not essential for adult viability. Targeting gene expression specifically in male germ cells allows one to study effects on cytokinesis in an otherwise healthy adult fly. Lastly, techniques are now available for culturing spermatocytes removed from larval to adult stage Drosophila, providing an opportunity to test the effects of genetic or chemical disruptions in real time (Brill et al., 2000; Inoue et al., 2004; Rebollo and Gonzalez, 2004; Wong et al., 2005).

Mutations in a large number of genes result in meiotic cytokinesis defects (Table 1.1). These mutations affect different stages of cytokinesis and the genes encode many of the proteins discussed in Section 1.1.2. Genes involved in general regulation of cytokinesis include polo, cdc37, australin (a testis-specific homologue of borealin-related (borr), part of the chromosomal passenger complex that includes Aurora B) and pbl (Carmena et al., 1998; Lange et al., 2002; Giansanti et al., 2004; Gao et al., 2008). Genes encoding microtubule binding proteins involved in spindle stability or signalling include fascetto (feo), orbit/mast and the kinesins KLP3A and KLP67A (Williams et al., 1995; Giansanti et al., 1998; Gandhi et al., 2004; Inoue et al., 2004; Verni et al., 2004; Gatt et al., 2005). Genes involved in actomyosin ring assembly, contraction or disassembly include dia, chic, tsr, sqh and citron kinase (Castrillon and

Wasserman, 1994; Gunsalus et al., 1995; Giansanti et al., 1998; Giansanti et al., 1999; Giansanti et al., 2001; Giansanti et al., 2004; Naim et al., 2004). Finally, genes involved in phospholipid regulation and/or membrane trafficking include fwd, giotto/vibrator (gio), bond, pex family genes, des-1, fws, (brunelleschi) bru, rab11, arf6 and syntaxin 5 (syx5) (Basu and Li, 1998; Brill et al., 2000; Xu et al., 2002; Farkas et al., 2003; Giansanti et al., 2004; Gatt and Glover, 2006; Giansanti et al., 2006; Dyer et al., 2007; Giansanti et al., 2007; Szafer-Glusman et al., 2008; Polevoy et al., 2009; Chen et al., 2010). Anillin is known to localize to cleavage furrows and ring canals in male germ cells (Hime et al., 1996; Giansanti et al., 1999). However, whether it plays a role in spermatocyte cytokinesis was previously unknown.

Table 1. Genes with roles in male meiotic cytokinesis Gene name Protein function Reference Regulation australin (aust) Member of the chromosomal passenger (Gao et al., 2008) complex, borealin-related homologue cdc37 Regulates Aurora B stability (Lange et al., 2002) pebble (pbl) GEF for RhoA (Giansanti et al., 2004) polo Polo kinase (Carmena et al., 1998) MT associated fascetto (feo) MT plus-end binding, central spindle (Verni et al., 2004) organization KLP3A Kinesin, central spindle organization (Williams et al., 1995; Giansanti et al., 1998) KLP67A Kinesin, formation and stability of central (Gandhi et al., 2004; Gatt et al., spindle 2005) orbit/mast MT plus-end binding, central spindle (Inoue et al., 2004) organization Actin cytoskeleton chickadee (chic) Profilin, promotes F-actin polymerization (Giansanti et al., 1998) citron kinase/ sticky Myosin II regulatory light chain kinase (Naim et al., 2004) (sti) diaphanous (dia) Formin, nucleates unbranched F-actin filaments (Castrillon and Wasserman, 1994; Giansanti et al., 1998; Giansanti et al., 2004) spaghetti squash (sqh) Myosin II regulatory light chain (Giansanti et al., 2001) twinstar (tsr) Cofilin, F-actin disassembly (Gunsalus et al., 1995; Giansanti et al., 1999) Lipid regulation bond Very-long-chain fatty acid synthesis (Giansanti et al., 2004; Szafer- Glusman et al., 2008) des-1/ infertile Sphingolipid delta-4 desaturase (Basu and Li, 1998) crescent (ifc) four wheel drive (fwd) Phosphatidylinositol 4-kinase III β (Brill et al., 2000; Giansanti et al., 2004; Polevoy et al., 2009) giotto/vibrator (gio) Phosphatidylinositol transfer protein (PITP) (Giansanti et al., 2004; Gatt and Glover, 2006; Giansanti et al., 2006) peroxins Peroxisome biogenesis, very-long-chain fatty (Chen et al., 2010) (pex1,2,10,12,13,16) acid synthesis Trafficking arf6 Regulates recycling and endocytosis (Dyer et al., 2007) brunelleschi (bru) Subunit of Golgi-localized TRAPII complex, (Giansanti et al., 2004) Trs120p orthologue four way stop (fws) Subunit of Golgi-localized COG complex, (Farkas et al., 2003; Giansanti et al., Cog5 orthologue 2004) rab11 Recycling endosome regulator (Giansanti et al., 2007; Polevoy et al., 2009) syntaxin 5 (syx5) Golgi-localized SNARE protein (Xu et al., 2002)

1.3.3 Individualization

Individualization occurs after meiosis and spermatid elongation and describes a process whereby a syncytial cyst of sixty-four elongated Drosophila spermatids is transformed into sixty-four independent spermatozoa (Figure 1.3B) (Tokuyasu et al., 1972; Fuller, 1993). Individualization begins with the formation of actin-based structures surrounding spermatid nuclei. As these coalesce, they become cone shaped and are termed investment cones. These structures traverse the length of the spermatid cyst, removing excess cytoplasm and organelles and investing each spermatid in its own membrane.

Investment cones are composed of two highly organized and separable arrays of F-actin – a dense meshwork at the front, requiring Arp2/3 for assembly, and a region of parallel bundles toward the rear, requiring profilin (Noguchi et al., 2006; Noguchi et al., 2008). Two myosin motors are required for spermatid individualization and appear to have non-overlapping roles. Myosin V is required for initial formation of investment cones, whereas myosin VI is required for investment cone movement (Hicks et al., 1999; Mermall et al., 2005). Myosin VI localizes to the leading edge of investment cones and is required for recruitment of cortactin and Arp2/3 (Rogat and Miller, 2002). The role of myosin VI in investment cone movement is proposed to be in tethering F-actin rather than transporting cargo, based on FRAP experiments that reveal a stable association with F-actin during movement (Noguchi et al., 2006).

Spermatid individualization provides an interesting biological context in which to investigate processes such as actin dynamics and membrane remodeling. It is a developmental system that still contains many unanswered questions. For example, what other proteins localize to actin-rich investment cones and how might they contribute to cone formation or progression? One protein that localizes to the investment cones in a similar manner to myosin VI turns out to be anillin (see below). Perhaps anillin acts as a scaffold in this context, linking F-actin to additional regulatory and structural molecules. Such a finding would broaden our understanding of Drosophila spermatogenesis in particular, and actin-based cell morphogenesis in general.

1.4 Thesis rationale and summary

Anillin is a conserved and essential scaffolding protein that plays a crucial role in cytokinesis. In the Drosophila testis, anillin localizes to several actin rich structures, including, but not limited

33 to, cleavage furrows of dividing cells. Defining specific functions for anillin outside of cytokinesis would broaden our understanding of this protein and the processes to which it contributes. In terms of cytokinesis, much remains to be deciphered. By studying anillin function, one can get at the basic properties and mechanics of cells undergoing cytokinesis, for example, the stability or dynamics of components of the contractile ring; the nature of the cytoskeletal-membrane connections; and the precise role of anillin in this process. In Chapter 3, plus the Appendix, I will present data that address these questions and further our understanding of the role of anillin in cytokinesis.

In Chapter 3, I begin with a description of anillin localization in developing male germ cells. This is followed by analysis of anillin phenotypes, specifically those involved in cytokinesis and in individualization. Because the individualization phenotype may be a secondary consequence of prior defects in cytokinesis, I continue with further analysis of the primary cytokinesis defect. I establish that anillin, the septin Sep2 and myosin II are all stably associated with the cleavage furrow. When anillin is depleted, septins are delocalized, and F- actin, myosin II and Rho are no longer restricted to the cleavage furrow. Live imaging reveals that both myosin and F-actin cycle around the membrane in the absence of anillin.

The latter part of Chapter 3 begins with the unexpected observation that expression of Drosophila E-cadherin fused to GFP (DE-cad-GFP) suppresses the anillin cytokinesis phenotype. Further examination of this finding reveals that DE-cad-GFP recruits armadillo and α-catenin, proteins normally absent in primary spermatocytes, to the cleavage furrow in dividing cells. Moreover, in the presence of DE-cad-GFP, F-actin is stabilized at the furrow, leading to a model in which adherens junction proteins can partially substitute for anillin and septins in linking F-actin to the overlying membrane during cytokinesis.

Finally, in the Appendix, I present additional data focused on the regulation of anillin in Drosophila male germ cells. Here, I begin with the finding that a mutation in the cell cycle regulatory gene, myt1, affects both subcellular localization of anillin and mobility of anillin on SDS-PAGE. I show that anillin is normally phosphorylated and the amount of phosphorylation depends, directly or indirectly, on dMyt1. Colocalization experiments suggest that anillin may be phosphorylated – or tightly associated with a protein that is phosphorylated – on tyrosine

34 residue(s). The implications of these findings are discussed and placed in the larger context of cytoskeletal biology and animal cell cytokinesis.

Chapter 2

Materials and Methods

2 Materials and Methods 2.1 Molecular biology

Molecular cloning was performed using standard techniques (Sambrook et al., 1989). Restriction enzymes and T4 DNA ligase were from New England Biolabs (NEB). Polymerase chain reaction (PCR) was performed using Phusion DNA Polymerase (Finnzymes, NEB) on an MJ Research PTC-200 PCR machine.

Plasmids for generating transgenic flies were made in the P element transformation vector tv3, which contains the spermatocyte-specific β2-tubulin (β2t) promoter and SV40 3’ sequences (Wong et al., 2005), or in pCaSpeR-tub::mCherry, a modified version of pCaSpeR4

(Pirrotta, 1988) containing the α1-tubulin promoter (generously provided by H. Krämer) (Marois et al., 2006) fused to mCherry (generously provided by R. Tsien) (Shaner et al., 2004) (J. Burgess and J. A. B., unpublished). A snapback RNA interference (RNAi) construct (Kalidas and Smith, 2002) directed against anillin, tv3::anillin-RNAi, was generated as an antiparallel genomic DNA–cDNA fusion, using the genomic BAC clone BACR04H12 (BACPAC Resources, Children’s Hospital Oakland Research Institute, Oakland, CA) and the anillin cDNA LD23793 (Berkeley Drosophila Genome Project, DGC Release 1) as templates for PCR amplification of 690 bp (genomic) or 530 bp (cDNA) encoding part of exon7 through exon 9. Genomic sequences were amplified as an EcoRI-SfiI fragment by PCR, using primers 5’- CGGAATTCTGAGGACAGTAGCGCCGGCATG-3’ and 5’- GCGGCCAAGATGGCCCTATAAACAACGAGT-3’. cDNA sequences were amplified as an SfiI-XbaI fragment using primers 5’- GCGGCCATCTAGGCCGTTAGCGCTCTGTTG-3’ and 5’- GCTCTAGATGAGGACAGTAGCGCCGGCATG-3’.

tv3::mRFP-anillin and tv3::GFP-anillin were made by fusing mRFP1 (a gift from R. Tsien) (Campbell et al., 2002) or monomeric EGFP (courtesy of E. Snapp) (Zacharias et al., 2002) to the amino terminus of the full-length anillin cDNA LD23793, which was amplified as an XmaI-KpnI fragment, using primers 5’- GCCCCGGGATGGACCCGTTTACTCAGCACA-3’ and 5’- GCGGTACCTCAGTGGGTGGTTCCCCAGGCG-3’, and ligated into pBluescript SK+

37 containing the EGFP or mRFP1 sequence. The fused sequences were transferred to tv3 as NotI- KpnI fragments. The calponin homology domain of utrophin (Utr-CH) was amplified from GFP- Utr-CH (kindly provided by B. Burkel and W. Bement) (Burkel et al., 2007) as an XhoI-SpeI fragment using primers 5’- GCCTCGAGATGGCCAAGTATGGAGAACATG-3’ and 5’- GCACTAGTTTAGTCTATGGTGACTTGCTGA-3’ and used to generate pCaSpeR- tub::mCherry-Utr-CH. All constructs were confirmed by DNA sequencing (The Centre for Applied Genomics, The Hospital for Sick Children).

2.2 Generation of anti-anillin antibody

Polyclonal antibodies were raised against the N-terminal portion of anillin as previously described (Field and Alberts, 1995), with the exception that the corresponding sequence of cDNA LD23793 encodes an additional 36 amino acids. DNA sequences encoding the N- terminal 409 amino acids of anillin were amplified by PCR as an EcoRI-NotI fragment using primers 5’- CGGAATTCATGGACCCGTTTACTCAG-3’ and 5’- GGGCGGCCGCGGGAGCAGATCCTATTGG-3’ and cloned into pGEX-4T-1 (GE Healthcare).

The resulting GST fusion protein (GST-anil1-409) was expressed in BL21[DE3] bacterial cells. Transformed cells were grown in liquid culture to an optical density (OD600) of ~0.6, induced with 0.1 mM IPTG and allowed to grow for another 3 hours. The culture was centrifuged at 10,000 g for 15 min at 4˚C. The supernatant was removed and the pellet was resuspended in 5ml BugBuster (Novagen, EMD4Biosciences) (containing 1:10,000 DNase and 1:10,000 RNase) per gram of wet cell paste. The solution was shaken for 15 min at room temperature and centrifuged at 16,000 g for 20 min at 4˚C. The supernatant was transferred to Poly-Prep Chromatography Columns (BioRad) containing 1 ml of equilibrated glutathione beads (GE Healthcare Glutathione Sepharose 4B) (1 ml of beads has a binding capacity of 5 mg GST) and mixed overnight at 4˚C. The columns were washed 3 times with 10 bed volumes (bv) of PBS (pH 7.2) and protein was eluted with elution buffer including glutathione (Sigma) (10 mM glutathione, 50 mM Tris, pH 8.0). The eluate was added to an equilibrated Nap-10 column to deplete glutathione and collected in 1.5 ml PBS. The concentration of the purified GST-anil1-409 antigen was measured using the Bradford assay.

Two specific pathogen free rabbits (#4091 and #4092) were used to generate antibodies. Preimmune serum was collected before injection of the antigen. A first bleed was collected after two initial injections, followed by a second bleed after one more injection. Animals were then exsanguinated and crude serum was isolated from whole blood. Blood was incubated at 37˚C for 4 hours (to promote clotting) and centrifuged in a table-top centrifuge for 20 min at 4˚C. The supernatant was removed, spun for an additional 10 min, removed again and stored for further use.

Total immunoglobulin-G (IgG) content was purified from crude serum. Protein A beads (BioRad Affi-Gel Protein A Gel) in 50% slurry were added to Poly-Prep columns and washed with 10 bv binding buffer (10 mM sodium phosphate, 0.15 M NaCl, pH 8.2). 1 ml of beads has a binding capacity of ~18mg IgG and crude serum generally contains ~10mg IgG/ml. Crude serum was mixed 1:1 with binding buffer, added to the capped column and mixed overnight at 4˚C. The column was drained and washed with 10 bv binding buffer. IgG was eluted in 1 ml fractions with 0.1 M sodium citrate (pH 3.0) and neutralized immediately with 200 µl/ml Tris (pH 9.0). 20 µl from each fraction was run on a 10% SDS-PAGE gel and the gel was stained with Coomassie Blue (R-250, BioRad). All fractions containing significant amounts of IgG were combined and used for further purification.

GST and GST-anil1-409 columns were constructed for antibody purification. Both GST and GST-anil1-409 (<5 mg) were purified as described above and added to Affigel-10 columns (BioRad Affi-Gel 10) containing 1 ml of equilibrated beads. The columns were capped and rotated for 4 hours at 4˚C. 0.1 ml 1 M ethanolamine (blocking agent) was added and the columns were rotated for 1 additional hour. Columns were drained, washed with 10 ml PBS (pH 7.2) and stored in 5 ml PBS. Before using, columns were washed with acid and base to remove any non-covalently linked molecules. Columns were washed with 10 bv 10 mM Tris (pH 7.5), 10 bv 100 mM glycine (pH 2.5), 10 bv 10 mM Tris (pH 8.8), 10 bv 100 mM triethylamine (pH 11.5) and 10 bv 10 mM Tris (pH 7.5).

Anti-anillin antibody was purified from total IgG with GST and GST-anil1-409 columns. The IgG solution was passed over the GST column twice to remove GST-specific antibodies. Collected flow-through was added to the GST-anil1-409 column, which was capped and rotated overnight at 4˚C. The column was washed with 20 bv 10 mM Tris (pH 7.5) and 20 bv 500 mM

NaCl, 10 mM Tris (pH 7.5). Acid elution was performed with 10 bv 100 mM glycine (pH 2.5). 1 ml fractions were collected and immediately neutralized with 200 µl 1 M Tris (pH 8.0). The column was washed with 10 bv 100 mM Tris (pH 7.5). Base elution was performed with 10 bv 100 mM triethylamine (pH 11.0). 1 ml fractions were collected and immediately neutralized with 200 µl HEPES (pH 6.0). 20 µl from each fraction of acid or base elution was run on a 10% SDS-PAGE gel and the gel was stained with Coomassie Blue. Anti-anillin antibody was detected only in acid elution fractions. All fractions containing significant amounts of IgG were mixed, aliquoted and stored at -20˚C or -80˚C.

Crude serum and purified anti-anillin antibody #4091 and #4092 were compared by immunofluorescence and immunoblotting. Purification reduced background for both methods. By immunofluorescence, no significant differences in staining were detected for either of the crude sera or purified antibodies. By immunoblotting, although crude serum recognized full- length anillin at similar levels compared to background for both antibodies, purified antibody #4092 appears to have a much lower titer than purified antibody #4091. For this reason, antibody #4091 was chosen for large-scale purification and experimental use.

2.3 Drosophila genetics

Flies were raised on standard cornmeal molasses agar at 25°C (Ashburner, 1990). Transgenes were introduced by injection of w1118 embryos as described (Wong et al., 2005). Stocks used in the generation of male germ cell clones were obtained from the Bloomington Drosophila Stock Center (Bloomington, IN): P{neoFRT}42D; ry605, w1118; P{neoFRT}42D P{Ubi- GFP(S65T)NLS}2R /CyO, y1 w1118 P{70FLP}3F/Dp(1;Y)y+; nocSco/SM6a, and P{PZ}anil03427,cn1/CyO; ry506, a P{PZ} insertion 53bp upstream of the start codon (anilPZ) (Doberstein et al., 1997; Field et al., 2005a). To make clones, flies were heat-shocked at 37°C for 1 hour daily until pupariation. UAS::anillin-RNAi flies were from the Vienna Drosophila RNAi Collection (VDRC #33465). Fly stocks were as follows: Bam-GAL4 (gift of D. McKearin) (Chen and McKearin, 2003); Sep2-GFP (gift of K. Hales, Davidson College) (Silverman-Gavrila et al., 2008); ubiquitin::DE-cad-GFP (provided by H. Oda via U. Tepass, University of Toronto) (Oda and Tsukita, 2001); Sqh-GFP (gift of R. Karess) (Royou et al.,

2002); β2t::CLC-GFP (gift of H. Chang, Purdue University). Flies expressing a secreted GFP

(Pfeiffer et al., 2000) under control of the β2t promoter (Wilson et al., 2006) have been described

40 elsewhere (Polevoy et al., 2009). GFPNLS flies used as controls in the DE-cad-GFP rescue experiment were the same as those used in generating germ cell clones (see above). fwd3/TM6B and Df(3L)7C/TM6B were previously described (Brill et al., 2000). w1118 flies were used as wild-type controls.

2.4 Fluorescence microscopy, imaging and analysis

Immunofluorescence was performed essentially as described (Hime et al., 1996), except that testis isolation buffer (TIB) (Casal et al., 1990) was used instead of TB-1. Unless specified, samples were fixed in PBS (pH 7.2) with 4% paraformaldehyde followed by permeabilization in PBS with 0.3% Triton X-100 and 0.3% sodium deoxycholate. For anti-DE-cad, anti-armadillo and anti-α-catenin, salt-free phosphate buffer (PB) (pH 7.4) was used instead of PBS and permeabilization was performed in PB plus 0.3% Triton X-100 (Niewiadomska et al., 1999). Antibodies used were as follows: 1:100 rabbit anti-anillin (see above), 1:20 mouse anti-myosin VI (3C7, a gift of K. Miller) (Kellerman and Miller, 1992), 1:150 mouse anti-peanut (4C9H4, Developmental Studies Hybridoma Bank (DSHB), Iowa City, IA) (Neufeld and Rubin, 1994), 1:100 mouse anti-Rho (p1D9, DSHB) (Magie et al., 2002), 1:50 rat anti-DE-cad (gift of U. Tepass) (Oda et al., 1994), 1:50 mouse anti-armadillo (N2 7A1, DSHB) (Riggleman et al., 1990) and 1:5000 guinea pig anti-α-catenin (gift of R. Sarpal and U. Tepass, Department of Cell and Systems Biology, University of Toronto). Rhodamine phalloidin (20 units/mL) and DAPI (5µg/mL final concentration) were used as recommended by the manufacturer (Molecular Probes).

To image live squashed preparations of Drosophila male germ cells expressing fluorescent fusion proteins (Sqh-GFP, mCherry-Utr-CH), testes were dissected in TIB, transferred to a microscope slide and cut with tungsten needles in TIB containing 8.3 µg/ml Hoechst 33342 (Sigma) to stain DNA. Samples were squashed with a coverslip before viewing. Live images of dividing spermatocytes prepared in fibrin clots (Wong et al., 2005) were acquired at 20-second intervals. Time zero in the figures was arbitrarily set at an equivalent stage of cytokinesis across all genotypes.

Phase-contrast and fluorescence images were acquired with a Zeiss Axiocam CCD camera on an upright Zeiss Axioplan 2 microscope using Axiovision software (Zeiss). Images of

41 separate fluorochromes from multiply stained tissues were collected individually and combined using Adobe Photoshop. When necessary, images were adjusted only for brightness and contrast. Unless otherwise stated, in cases in which direct comparison of images was required, images were acquired using identical exposure times and adjusted in an identical manner.

2.5 FRAP image acquisition and data analysis

Fluorescence recovery after photobleaching (FRAP) was performed on live cells prepared as described (Wong et al., 2005), using a Zeiss Axiovert 200 equipped with a Hamamatsu C9100- 13 EM-CCD camera, Yokogawa spinning disk confocal scan head, diode-pumped solid state laser lines (Spectral Applied Research: 405nm, 491nm, 561nm, 638nm) and a Ludl motorized XY stage. Images were acquired with a 63X/1.3 Zeiss Plan Apo water-immersion objective, with an additional 1.5x magnification lens in front of the camera. Photobleaching was performed using the Photonic Instruments Mosaic FRAP illuminator (488nm). Acquisition and analysis were performed using Improvision Volocity 5 (PerkinElmer).

Bleaching was performed on one visible edge of the cleavage furrow, where the opposite edge was used as an internal, unbleached control. Volocity FRAP analysis software was used to measure the t1/2 values for the PLCδ-PH-GFP marker. GFP images were corrected for photobleaching in Volocity. Circular regions of interest of 2 µm diameter were centered on the visible edges of cleavage furrows for bleached and unbleached sides at each time point. Mean fluorescence intensity values were measured in Volocity and transferred to Microsoft Excel. Fluorescence intensity values were normalized using the equation: (yx – ymin)/(ymax – ymin), where y is mean fluorescence intensity and yx is the value of y at a given timepoint (x). Graphs were generated in Microsoft Excel. For GFP-anil, Sep2-GFP and Sqh-GFP, percentage recovery was calculated by taking the mean fluorescence intensity value of 6 individual values centered on 5 minutes postbleach, normalizing against unbleached values, and dividing by the prebleached value.

2.6 Immunoblotting

For testis immunoblots, 20-30 pairs of testes per genotype were dissected in TIB and transferred directly to eppendorf tubes containing SDS-PAGE sample buffer. Samples were pipetted up and down to dissolve tissue, boiled for 5 minutes and centrifuged at maximum speed for 1 minute.

Proteins were separated on 8,10 or 12% SDS polyacrylamide gels and transferred to Amersham Hybond-ECL nitrocellulose membranes (GE Healthcare) using a Hoefer miniVE system (Hoefer, Inc.) Blocking and antibody incubations were in Tris-buffered saline with 0.05% Tween-20 (TBST) containing 4% nonfat dry milk (Bio-Rad Blotting Grade Blocker 170-6404). Washes were in TBST. Primary antibodies were used at the following dilutions: anti-anillin (1:1000), anti-armadillo (1:500), anti-α-catenin (1:5000), anti-β-tubulin N-357 (Amersham, GE Healthcare; 1:4000). HRP-conjugated secondary antibodies (Jackson ImmunoResearch Laboratories, Inc.) were used at 1:10,000. Signals were detected using Amersham ECL Plus Western Blotting Detection Reagents.

Chapter 3

Results

The majority of the work described in this chapter was published in May 2010, Molecular Biology of the Cell, 21 (9): 1482-93 (Copyright 2010 by The American Society for Cell Biology; http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E09-08-0714), with the following list of authors: Philip Goldbach, Raymond Wong, Nolan Beise, Ritu Sarpal, William S. Trimble and Julie A. Brill. Unless otherwise indicated, I performed all experiments described herein. Live imaging experiments were carried out in collaboration with Raymond Wong. I generated the transgenic fly lines and set up crosses. Raymond prepared the larval clot preps. I captured images for the FRAP experiments and Raymond captured other real-time images. Mouse L- fibroblast experiments were performed by Nolan Beise and analyzed by Nolan Beise and William S. Trimble.

3 Results 3.1 Introduction

Cytokinesis has long been recognized to involve formation and constriction of a contractile ring composed of F-actin and non-muscle myosin II (Satterwhite and Pollard, 1992; Rappaport, 1996). Bipolar filaments of myosin II are thought to draw F-actin together in a purse-string like manner to constrict the contractile ring. Studies in yeast and mammalian cells suggest that actin and myosin turn over during constriction, leading to an idea that F-actin and myosin are constantly recruited and disassembled at the cleavage furrow during cytokinesis (Pelham and Chang, 2002; Murthy and Wadsworth, 2005). However, recent work in the early C. elegans embryo suggests that actin and myosin do not turn over, but rather disassemble over time (Carvalho et al., 2009). In either case, a tight link must be established between the actomyosin ring and the plasma membrane at the equator, and this attachment must be maintained during late stages of cytokinesis.

Anillin was originally identified in Drosophila melanogaster embryo extracts as an actin binding and bundling protein that localizes to the cleavage furrow during cytokinesis (Miller et al., 1989; Field and Alberts, 1995). Subsequent studies revealed anillin to be a conserved multidomain scaffolding protein with an essential role in cytokinesis (Section 1.2). Notably, since many of the published experiments describing anillin function have been carried out in tissue culture models, the mechanism by which anillin promotes conventional cytokinesis in a multicellular organism remains poorly understood.

Although anillin is widely expressed during development, most functional studies have so far been carried out in tissue culture cells in vitro. In the Drosophila testis, anillin localizes to a number of different structures in developing male germ cells (Hime et al., 1996; Giansanti et al., 1999). Similar to other cell types (reviewed in Section 1.2.2), anillin localizes to nuclei in premeiotic cells that are in interphase. In mitotic or meiotic cells undergoing cytokinesis, it is tightly associated with the cleavage furrow. After cytokinesis, anillin remains associated with mitotic and meiotic ring canals and colocalizes with at least three septins – peanut, Sep1 and Sep2 (Hime et al., 1996). This marks a difference between ring canals in males and females;

45 during oogenesis, ring canals become enriched in F-actin and actin-associated proteins (not found in males) and lose anillin (Robinson et al., 1994; Field and Alberts, 1995). Anillin and septins are proposed to provide structural support to male germline ring canals in the absence of F-actin (Hime et al., 1996). In addition to its localization to cleavage furrows and ring canals, we describe anillin in other post-meiotic structures (see below). Importantly, although anillin localization had been well characterized in developing male germ cells, its function during sperm development was not previously established.

In the first part of this chapter, I present data showing a requirement for anillin in meiotic cytokinesis. In anillin-depleted spermatocytes myosin II and Rho are no longer tightly associated with the cleavage furrow and septins are completely absent. I conclude that anillin plays a role in anchoring the CR to the furrow membrane. This does not preclude an additional role for anillin in regulating membrane trafficking. Indeed, many other genes that are required in late stages of cytokinesis affect membrane trafficking. In the second part of this chapter, I explore the effect of anillin depletion on membrane trafficking. Although no defects are detected, expression of the trafficking marker DE-cad-GFP turns out to suppress the cytokinesis defects caused by loss of anillin.

DE-cadherin is a Drosophila homologue of classical cadherins such as mammalian epithelial cadherin (E-cadherin). Adherens junction complexes, which contain E-cadherin and associated catenins, are able to link the plasma membrane to the actin cytoskeleton (reviewed in Section 1.1.3). In the Drosophila testis, DE-cadherin function has been characterized within the apical hub – the site of the niche that maintains germline and somatic stem cells (Section 1.3.1). DE-cadherin is required for maintenance of both germline and somatic stem cells (Yamashita et al., 2003; Wang et al., 2006; Boyle et al., 2007; Voog et al., 2008). Homophilic interactions between DE-cadherin in the membranes of stem cells and adjacent hub cells is proposed to keep stem cells in contact with self-renewing factors (Wang et al., 2006; Voog et al., 2008). Additionally, DE-cadherin is thought to play an active role in maintaining stem cell polarity through direct or indirect association with adenomatous polyposis coli suppressor protein (APC) (Yamashita et al., 2003). To our knowledge, DE-cadherin localization and function have not previously been tested in later stages of male germ cell development.

3.2 Results

3.2.1 Anillin is found in a variety of actin-associated structures during spermatogenesis

Anillin was previously reported to localize to contractile rings and ring canals during spermatogenesis (Hime et al., 1996; Giansanti et al., 1999). At the onset of this study, the function of anillin in these structures was poorly understood. In addition, immunostaining of whole testes with a scarce anti-anillin antibody (Field et al., 1995) revealed additional uncharacterized areas of anillin localization (J. A. Brill, unpublished).

To further characterize anillin localization, I generated a new polyclonal antibody using an epitope similar to that previously described by Field et al. (1995; see Section 2.2). I tested crude antisera raised in parallel in two rabbits, #4091 and #4092, in immunoblotting and immunofluorescence experiments. Both antisera recognized a protein at the expected molecular weight for anillin of ~190 kDa by SDS-PAGE (Figure 3.1C). Testis lysates contained significantly more of this protein than lysates from whole adult flies. In addition, antisera recognized a unique band in lysates from testes expressing a GFP-anillin construct (see below) corresponding to the increased molecular weight of the tagged protein. By immunofluorescence both antisera recognized protein in cleavage furrows and ring canals (see below). Because of apparent non-specific binding suggested by immunoblotting and immunofluorescence, I decided to affinity purify the antisera (see Materials and Methods) (Figure 3.1D and E). Both antibodies exhibited less non-specific binding after affinity purification. In immunoblotting of testis lysates, purified antibody #4091 exhibited a higher titer against full-length anillin than antibody #4092 (Figure 3.1F). For this reason I chose to use antibody #4091 (hereafter called anti-anillin antibody) for subsequent experiments.

Immunostaining of developing male germ cells with anti-anillin antibody revealed anillin localization to mitotic ring canals (Figure 3.2A), contractile rings (Figure 3.2B) and ring canals at the growing ends of elongating spermatid cysts (Figure 3.2C and D). In addition, anillin localized to the dense body, an actin- and tubulin-rich structure that associates with haploid nuclei during nuclear shaping (Figure 3.2D and E). During investment cone formation, anillin was found in puncta near the investment cones (Figure 3.2F). In cysts in which the investment cones started to move away from the nuclei, anillin puncta appeared to coalesce (Figure 3.2G),

47 becoming highly concentrated at the leading edge of mature cones as they traversed the length of individualizing cysts (Figure 3.2H). Anti-anillin antibody staining was specific, as the signal was largely absent from male germ cells depleted for anillin (see below).

Anillin localization was further confirmed by generation and examination of fluorescently-tagged anillin fusion proteins. Specifically, I generated N-terminal GFP-anillin and monomeric red fluorescent protein- (RFP-) anillin constructs in a Drosophila transformation vector containing the testis-specific β2-tubulin promoter. I injected embryos and selected transgenic flies, which I used to examine localization of the fluorescent fusion proteins in the testis. Localization of GFP-anillin and RFP-anillin resembled that seen by immunostaining (Figure 3.3). GFP-anillin and RFP-anillin localized to nuclei, cleavage furrows and ring canals (Figure 3.3A-C). Interestingly, mitotic ring canals contained small amounts of GFP-anillin, suggesting that there is some turnover of anillin in ring canals (Figure 3.3A). Alternatively, some GFP-anillin may be expressed prior to mitotic divisions, although this is unlikely, as the

β2-tubulin promoter is active only in mature primary spermatocytes (Michiels et al., 1989; Hoyle and Raff, 1990). GFP-anillin, which produced a brighter signal than RFP-anillin, also localized to later developmental structures, including dense bodies and puncta just distal to elongating nuclei (Figure 3.3D). Thus, it is clear from immunostaining experiments and localization of tagged proteins that anillin is highly concentrated in several actin-rich structures in developing male germ cells.

Figure 3.1. Generation of anti-anillin antibody.

(A,D,E) Polyacrylamide gels stained with Coomassie Blue and (B,C,F) immunoblots. (A) The purified antigen, GST-anil1-409, migrates slightly above the predicted molecular weight of ~70 kDa and appears to be partially degraded. GST migrates at ~30 kDa. (B) Preimmune sera and crude antisera (C) were tested for ability to recognize anillin and for antigen specificity (C). (B) Preimmune serum #4091 does not recognize a prominent protein species in wild-type testis lysates at the expected molecular weight for full-length anillin (~190 kDa). Dilution, 1:200. (C) Lysates from male and female adult flies, wild-type testes and testes from flies expressing GFP- anillin were probed with crude antiserum #4091, diluted 1:200. The antiserum recognizes a protein at ~190 kDa in all samples. The appearance of a slower migrating protein in lysates from GFP-anillin-expressing testes confirms that the antiserum recognizes anillin. (D-F) Affinity purification of anti-anillin antibody. In a two-step process, total IgG was purified from crude serum (D) and anti-anillin antibody was purified from total IgG (E). Protein at the expected molecular weight for IgG heavy chain (~50 kDa) is detected in a number of elution fractions at each step. (F) Wild-type testis lysates were probed with indicated dilutions of crude serum or purified anti-anillin antibody from rabbits #4091 and #4092. Note that purified antibody #4091 exhibits a higher titer against full-length anillin than purified antibody #4092.

Figure 3.2. Anillin localizes to actin-rich structures during spermatogenesis.

(A-H) Fluorescence micrographs of male germ cells stained for F-actin (red), anillin (green) and DNA (blue) and (A’-H’) corresponding grayscale images of anillin alone. (A) Primary spermatocytes with F-actin in the fusome and anillin in mitotic ring canals (arrows) and nuclei (arrowheads). (B) Dividing spermatocytes showing colocalization of F-actin and anillin in the cleavage furrow (arrows). (C) Cyst of 64 early elongating spermatids with anillin in ring canals (arrow). (D) Cyst of elongating spermatids showing a high concentration of F-actin near ring canals (arrow). Anillin and F-actin are also present in the dense body (arrowhead), which forms a crescent along one side of each spermatid nucleus. (E) Group of 64 elongated sperm nuclei. Anillin and actin persist in the dense body (arrow) throughout nuclear shaping. (F) Actin cones forming over three clusters of 64 nuclei. Anillin localizes to puncta (arrow) in the vicinity of the actin cones (arrowhead). (G) Puncta of anillin coalesce at the leading edge of actin cones (arrows) as they prepare to move away from the mature sperm nuclei. (H) Anillin localizes to the leading edge of actin cones (arrow) that have progressed along the length of the cyst. Bars (A-B) 20µm, (C-D) 20µm and (E-H) 20µm.

Figure 3.3. GFP- and RFP-anillin localize to same structures recognized by anti-anillin antibody.

(A-D) Fluorescence micrographs of male germ cells expressing GFP-anillin (A,B,D) or RFP- anillin (C) and corresponding phase-contrast micrographs. (A) GFP-anillin and RFP-anillin (not shown) localize to spermatocyte nuclei (arrows). (A’) When image brightness is increased, GFP-anillin is visible in mitotic ring canals (arrowheads). (B) GFP-anillin and RFP-anillin (C) localize to cleavage furrows (arrows) in dividing spermatocytes. (D) GFP-anillin also localizes to dense bodies (arrowhead) and in puncta located slightly caudally (arrow). These puncta are seen more clearly with increased image brightness (D’). Bars (A) 20µm and (B-D) 20µm.

3.2.2 Anillin is required for spermatocyte cytokinesis

To test whether anillin is required for male germ cell development, I used the FLP/FRT system (see Materials and Methods) to generate clones of male germ cells homozygous for a lethal P- element insertion in anillin (anilPZ). Unlike wild-type spermatids (Figure 3.4B), anilPZ mutant spermatids (Figure 3.4C) were multinucleate, indicating a defect in male meiotic cytokinesis (Romrell et al., 1972). To confirm that this was due to the anilPZ mutation, I expressed RFP- anillin, which fully rescued the cytokinesis defect of anilPZ mutant clones (Figure 3.4D).

To further explore the requirement for anillin in male germ cells, I used RNAi induced by expression of double-stranded RNA (dsRNA) directed against sequences from the anillin gene (see Section 2.1). Two different dsRNAs were used – one a genomic-cDNA hybrid RNAi construct expressed under control of the primary spermatocyte-specific β2-tubulin promoter genomic-cDNA (β2t>anillin-RNAi ) (Hoyle and Raff, 1990; Wong et al., 2005); the other a UAS- dsRNA construct obtained from the Vienna Drosophila RNAi Center (VDRC) (UAS>anillin- RNAiVDRC) and expressed in early primary spermatocytes using Bam-Gal4 (Chen and McKearin, 2003; Dietzl et al., 2007). These dsRNAs, which were directed against different portions of the anillin gene (Figure 3.4A), also caused formation of multinucleate cells (Figure 3.4F). Since expression of UAS>anillin-RNAiVDRC caused a highly penetrant cytokinesis defect, I used this line in subsequent experiments.

To confirm that anillin protein levels were knocked down, I examined dsRNA-expressing male germ cells using anti-anillin antibody. In wild-type spermatocytes, anillin localized to interphase nuclei and to mitotically formed ring canals (Figure 3.4G). In spermatocytes expressing dsRNA directed against anillin, anillin protein levels were greatly reduced in spermatocyte nuclei, but remained in mitotic ring canals, which formed prior to dsRNA expression (Figure 3.4H). In meiotically dividing wild-type male germ cells, anillin colocalized with F-actin in contractile rings (Figure 3.4I). However, in dsRNA-expressing cells, anillin was absent from cleavage furrows and F-actin was either diffusely localized or absent (Figure 3.4J, and see below). Immunoblotting of whole testis proteins confirmed that anillin levels were greatly reduced in dsRNA-expressing cells (Figure 3.4K). Thus, RNAi directed against anillin depleted anillin protein at post-mitotic stages.

Figure 3.4. Anillin is required for spermatocyte cytokinesis.

(A) Schematic of anillin gene sequence showing regions targeted by RNAi constructs. Grey boxes represent exons, connecting lines represent introns and thin green and red bands represent start and stop codons, respectively. UAS>anillin-RNAiVDRC (green box) and β2t>anillin- RNAigenomic-cDNA (blue boxes, genomic sequences; red box, reverse complement cDNA) target non-overlapping sequences. Drawing of anillin genomic region adapted from Flybase (http://flybase.org/reports/FBgn0261385.html). (B,C and E,F) Phase-contrast micrographs of early round spermatids, showing nuclei (white disks) and mitochondrial derivatives (dark organelles). (B,C) Spermatids from testes in which anillin mutant (anilPZ) clones were generated by FLP-FRT mediated recombination (see Materials and Methods). Wild-type spermatids are marked by nuclear GFP and have one nucleus per mitochondrial derivative (yellow arrowhead) (B). anillin mutant spermatids, marked by absence of GFP, contain multiple nuclei and enlarged

53 mitochondrial derivatives (yellow arrowheads), indicating failure of meiotic cytokinesis (C). (D,D’) Expression of RFP-anillin rescues the cytokinesis defects of anillin mutant clones. Fluorescence micrograph (D) of a GFP-negative cyst of rescued, mononucleate early spermatids (D’) expressing RFP-anillin (red). (E,F) Spermatids from wild-type (E) testes and testes in which male germ cells are expressing dsRNA targeted against anillin (F). Note that the samples in (D-F) were not flattened as much as those in (B,C). (G,H) Fluorescence micrographs of primary spermatocytes stained for anillin. (G’,H’) Corresponding merged images showing anillin (green) and F-actin (magenta). Anillin localizes to nuclei and mitotic ring canals in wild type (G), whereas in dsRNA-expressing cells, anillin is still present in mitotic ring canals but levels of nuclear anillin protein are greatly reduced (H). (I-I’’,J-J’’) Dividing spermatocytes stained for anillin (green), F-actin (red) and DNA (blue). In cleavage furrows (arrows) of wild- type spermatocytes, F-actin (I) and anillin (I’) colocalize (I’’), whereas in dsRNA-expressing cells F-actin appears diffuse (J) and anillin is absent (J’). Bars, 20µm. (K) Immunoblot showing reduced anillin (~190 kDa) levels in dsRNA-expressing male germ cells as compared to wild type. β-tubulin (~50 kDa) is used as a loading control.

I undertook two additional methods of disrupting anillin in a manner that would allow for examination of a testis phenotype in viable adults. Both of these methods had the advantage that they would allow for introduction of mutated anillin constructs to probe structure-function relationships. First, I generated flies expressing a wild-type copy of anillin under control of the Hsp70 heat-shock (hs) promoter (hs-anillin) (construct generated by J. A. Brill, unpublished). The aim of the experiment, based on a similar study of the septin peanut (Neufeld and Rubin, 1994), was to use a heat-shock regimen to provide flies homozygous for a strong anillin allele with enough wild-type protein to develop into adults. Once the adult stage is reached, flies are no longer heat-shocked, no new anillin is produced and a loss-of-function phenotype should present itself. I crossed the hs-anillin construct separately into flies homozygous for two strong anillin alleles: anilPZ, the allele used in generation of mutant clones, and anil8, a zygotic lethal allele previously described (Field et al., 2005a; O'Farrell and Kylsten, 2008). Despite varying the heat-shock regimen and confirming that anillin protein was being made by SDS-PAGE, I was unable to rescue the mutant flies to adulthood. One possible explanation for the lack of rescue could be that the developing organism is very sensitive to anillin levels and precise amounts are needed at the right time and in the right place. Another explanation could be a developmental requirement for additional anillin isoforms encoded by genomic sequences not present in the rescue construct, although this seems unlikely, as the three predicted anillin protein products are nearly identical (http://flybase.org). Alternatively, the mutant chromosomes could have picked up additional lethal mutations that cannot be rescued by anillin expression.

Second, I generated another RNAi construct by separating anillin 3’ UTR sequence and its reverse complement with an intron spacer (Lee and Carthew, 2003). This dsRNA would have the advantage of not targeting transgenic anillin constructs lacking the endogenous 3’ UTR. I examined 20 fly lines expressing the RNAi construct, either alone or in combination, and none showed testis phenotypes. The lack of phenotype may have been due to inadequate knockdown levels. Although I took as much sequence from the 3’ UTR as possible, the final sequence length was still shorter than recommended (Lee and Carthew, 2003). Specifically, my construct contained 136 nucleotides instead of the 500 to 1000 nucleotides recommended for effective knockdown.

Since generation of mutant clones and expression of two independent dsRNAs directed

55 against anillin proved effective in disrupting anillin function in the testis, I continued to use these tools to examine further the role of anillin in meiotic cytokinesis and to probe for additional roles in spermatogenesis.

3.2.3 Anillin knockdown causes defects in investment cone morphology and function

To determine if anillin has later functions in spermiogenesis, I examined dsRNA- expressing male germ cells for defects in differentiation and individualization. Loss of anillin had no obvious effect on polarity, elongation or differentiation of spermatid cysts. However, actin cone formation appeared somewhat aberrant. In male germ cell cysts expressing dsRNA directed against anillin, investment cones exhibited two phenotypes: some appeared thinner than normal and lacked myosin VI (Figure 3.5B), whereas others were short with diffuse myosin VI (Figure 3.5C). Similar investment cone defects were observed in male germ cells mutant for Fwd, which is also required for cytokinesis (Figure 3.5D and E), suggesting that failure to make normal investment cones may be a secondary consequence of the defect in cytokinesis. For this reason, I focused on the role of anillin in meiotic cytokinesis.

Figure 3.5. Anillin is required for normal investment cone morphology.

(A-E’’) Fluorescence micrographs of investment cones stained for F-actin (magenta) and myosin VI (green). In wild-type individualizing spermatids F-actin localizes throughout the cone (A) whereas myosin VI is tightly associated with the leading edge (A’). In cells expressing dsRNA directed against anillin (B-C’’) or in cells from fwd mutants (D-E’’), investment cones are either long and thin (B,D) with no detectable myosin VI (B’,D’) or compact (C,E) with diffuse myosin VI (C’,E’). Bar, 20µm.

3.2.4 Anillin, septins and myosin II are stably associated with the contractile ring

For anillin and septins to provide a stable anchor for the contractile ring, they should remain tightly associated with the cleavage furrow during cytokinesis. To test this idea, I examined GFP fusions to anillin and the septin Sep2 in FRAP experiments. I also tested a GFP fusion to myosin II regulatory light chain (Sqh), which was expected to be more dynamic during cleavage. Using live spermatocytes in fibrin clots (Royou et al., 2002; Wong et al., 2005; Wong et al., 2007), I photobleached one side of the cleavage furrow and measured fluorescence intensity levels in the bleached and unbleached sides over time. As a positive control, I examined the behavior of

PLCδ-PH-GFP, a marker for plasma membrane-associated PIP2 (Wong et al., 2005), which recovered rapidly with a t1/2 of 6.4 ± 1.3 seconds (Figure 3.6A,E; mean ± SD; n=5). In contrast, GFP-anillin (GFP-anil), Sep2-GFP and Sqh-GFP showed only low levels of recovery even after 5 minutes. Specifically, GFP-anil reached 37.2 ± 5.1% (Figure 3.6B,F; mean ± SD; n=5) of initial levels, Sep2-GFP reached 20.6 ± 3.9% (Figure 3.6C,G; mean ± SD; n=5) and Sqh-GFP reached 29.5 ± 8.1% (Figure 3.6D,H; mean ± SD; n=4). These numbers likely overestimate the degree of turnover of GFP-anil, Sep2-GFP and Sqh-GFP, as a significant proportion of the observed recovery may be due to unbleached material entering the region of interest during constriction (see also Carvalho et al., 2009). Thus, these data suggest that core components of the cleavage furrow and contractile ring exist in stable rather than dynamic structures during Drosophila spermatocyte cytokinesis.

Figure 3.6. Anillin, septins and myosin II are stably associated with the cleavage furrow.

Fluorescence micrographs (A-D) and corresponding graphs of dividing spermatocytes examined for FRAP (E-H). One example for each fluorescent marker is shown. (A-D) Micrographs show cells prior to bleaching (pre-bleach), immediately after bleaching (postbleach) and during recovery. The bleached area is shown (box with dotted lines). Times are in minutes: seconds. (E-H) Graphs show recovery of fluorescence in bleached area versus control (unbleached). Fluorescence intensity is indicated relative to starting intensity, which was set at 1. Time is in seconds. (A,E) PLCδ-PH-GFP, which associates with plasma membrane PIP2, recovers rapidly after photobleaching, whereas GFP-anillin (B,F), Sep2-GFP (C,G) and Sqh-GFP (D,H) fail to recover even after five minutes. Bar, 10µm.

3.2.5 Anillin recruits septins and maintains Rho, F-actin and myosin II at the cleavage furrow

Since anillin is known to bind septins, F-actin, myosin II and Rho, I examined localization of these proteins in dividing spermatocytes depleted for anillin. In wild-type spermatocytes, the septins peanut and Sep2 were concentrated in the cleavage furrow (Figure 3.7A and C), whereas septins were absent from the cleavage furrow in male germ cells expressing dsRNA directed against anillin (Figure 3.7B and D). F-actin was recruited to the cleavage furrow in anillin- depleted spermatocytes, but its localization appeared diffuse, especially at late stages of cytokinesis (compare Figure 3.7A’ and B’). Rho localization also appeared defective in dividing spermatocytes lacking anillin. Anillin colocalized with Rho at the cleavage furrow in wild-type cells (Figure 3.7E). However, in male germ cells depleted for anillin, the small amount of Rho detected at the cleavage furrow appeared diffuse (Figure 3.7F). As a control I examined septin and F-actin localization in testes expressing dsRNA directed against another gene, type II PI4K (PI4KII), which is dispensable for cytokinesis, but is required in late stages of sperm development (Jason Burgess, unpublished). Male germ cells expressing dsRNA against PI4KII exhibited normal peanut and F-actin localization (not shown), demonstrating that the anillin knockdown phenotype is not a non-specific result of expressing dsRNA.

To determine whether anillin is required to restrict myosin II to the equator of dividing spermatocytes, I examined live cells embedded in fibrin clots for localization of Sqh-GFP. In wild-type cells, Sqh-GFP was concentrated in the cleavage furrow during all stages of constriction (Figure 3.7G; n=3/3 cells). Sqh-GFP was no longer restricted to the equator in spermatocytes depleted of anillin. Rather, Sqh-GFP appeared to move around the periphery of the cell during cleavage (Figure 3.7H; n=5/6 cells). In addition, the membranes of dividing anillin-depleted cells exhibited blebbing and undulations not observed in wild type. Thus, anillin is required for localization of septins, and also for maintenance of Rho, F-actin and myosin II, at the cleavage furrow in dividing Drosophila spermatocytes.

To further examine Sqh-GFP behavior in anillin depleted cells, I performed FRAP on spermatocytes co-expressing Sqh-GFP and dsRNA targeted against anillin (Figure 3.8). In these cells, the bleached side of the furrow recovered at a greater rate and to a greater extent than in control cells (compare Figure 3.8A,B with Figure 3.6D,H). Specifically, fluorescence intensity

60 in the bleached region recovered to more than 90% of its original value after 185 ± 54.5 seconds (mean ± SD, n=5/6 cells). Meaningful half times of recovery could not be calculated since fluorescence recovery was not related to standardized rates of GFP molecule diffusion. Instead, delocalized Sqh-GFP flowed in and out of the furrow region, causing the bleached and unbleached sides of the furrow to be indistinguishable after a short period of time. Thus, anillin is essential for persistent association of myosin with the cleavage furrow.

Figure 3.7. Anillin is required for recruitment of septins and stabilization of Rho and myosin II at the cleavage furrow.

(A-A’’,B-B’’) Dividing spermatocytes stained for the septin peanut (Pnut) (green), F-actin (red) and DNA (blue). In cleavage furrows (arrows) of wild-type spermatocytes, peanut (A) and F- actin (A’) colocalize in cleavage furrows (A’’), whereas in dsRNA-expressing cells peanut is absent (B) and F-actin appears diffuse (B’). (C-D’) Dividing spermatocytes expressing Sep2- GFP (green) and stained for DNA (blue) with corresponding phase-contrast micrographs (C’’,D’’). Sep2-GFP localizes to cleavage furrows in wild-type cells (C), but is absent in dsRNA-expressing cells (D). (E-F’) Dividing spermatocytes stained for Rho (green), anillin (red) and DNA (blue) with corresponding phase contrast micrographs (E’’,F’’). In cleavage furrows of wild-type spermatocytes, Rho (E) and anillin colocalize in cleavage furrows (E’),

62 whereas in dsRNA-expressing cells Rho appears diffuse (F) and anillin is absent (F’). Bar, 20µm. (G,H) Time-lapse images showing the change in localization of Sqh-GFP (myosin regulatory light chain) during cytokinesis. In control cells, Sqh-GFP is tightly associated with the furrow (G), whereas in dsRNA-expressing cells Sqh-GFP moves around the cell cortex in an oscillatory manner (H). The images shown were taken at 40 sec intervals. Bar, 5µm.

Figure 3.8. Anillin is required for tight association of myosin II with the cleavage furrow.

Fluorescence micrograph (A) and corresponding graph (B) of a dividing spermatocyte, expressing Sqh-GFP and dsRNA directed against anillin, examined for FRAP. (A) Micrograph showing cell prior to bleaching, immediately after bleaching and during recovery. The bleached area is shown (box with dotted lines). Times are in minutes: seconds. (B) Graph showing recovery of fluorescence in bleached area versus control (unbleached). Fluorescence intensity is indicated relative to starting intensity, which was set at 1. (A,B) Fluorescence intensity increases and decreases in both bleached and unbleached areas as Sqh-GFP flows in and out of the furrow region. Time is in seconds. Bar, 10µm.

3.2.6 Ectopically expressed DE-cad-GFP as a marker for membrane trafficking in male germ cells

Since loss of anillin affects late telophase, a stage of cytokinesis when membrane trafficking is required, I tested the requirement for anillin in localization of several membrane trafficking markers. In early wild-type dividing spermatocytes, a GFP fusion to Drosophila E-cadherin (DE-cad-GFP) localized to the cell cortex and to puncta at the poles of the cells (Figure 3.9A). By mid-cytokinesis, DE-cad-GFP was found in puncta at the poles of the cells and also started to accumulate in puncta at the equator (Figure 3.9B). In late telophase, puncta of DE-cad-GFP were highly concentrated at the equator of dividing cells (Figure 3.9C). DE-cad-GFP localization appeared normal in spermatocytes depleted for anillin (Figure 3.9D). Similarly, in both wild-type and anillin-depleted spermatocytes, a GFP fusion to clathrin light chain (CLC- GFP) and a secreted GFP marker (sGFP) localized to the equator of dividing cells (Figure 3.10A- D). These data suggest that anillin does not play an important role in membrane trafficking to the cleavage furrow.

3.2.7 Loss of anillin is suppressed by expression of DE-cadherin

Remarkably, I found that expression of DE-cad-GFP, but not CLC-GFP or sGFP, had an unanticipated effect on cytokinesis in anillin-depleted cells. Male germ cells co-expressing DE- cad-GFP and dsRNA (UAS>anillin-RNAiVDRC; see Section 3.3.2) directed against anillin had an increased frequency of successful cytokinesis, relative to cells expressing dsRNA alone (Figure 3.9E). In wild type, 99.6% of spermatids were mononucleate, indicating successful meiotic cytokinesis. In contrast, only 4.4% of spermatids depleted of anillin were mononucleate. 16.7% of anillin-depleted spermatids had two nuclei per cell, indicating failure of cytokinesis during meiosis I or meiosis II, and 78.9% of spermatids had four nuclei per cell, indicating cytokinesis failure during both meiosis I and II. Expression of a nuclear GFP marker (GFPNLS) had no effect on spermatids depleted of anillin. However, ubiquitous expression of DE-cad-GFP resulted in significant suppression of the cytokinesis defect caused by anillin loss: 39.1% of spermatids were mononucleate, 19.3% were binucleate and 41.6% had four nuclei. Suppression of the cytokinesis defect by DE-cad-GFP was specific to anillin, as DE-cad-GFP had no effect on cytokinesis failures due to loss of fwd.

DE-cad-GFP expression also suppressed cytokinesis defects in male germ cells

genomic-cDNA expressing a different dsRNA directed against anillin, β2t>anillin-RNAi (Figure 3.9E; RNAi construct represented in Figure 3.4A). Two fly lines with independent chromosomal insertions of the construct were tested (RNAi#1 and RNAi#2). In flies expressing RNAi#1, 35.0% of spermatids were mononucleate, 31.0% were binucleate and 34.0% had four nuclei whereas in flies co-expressing RNAi#1 and DE-cad-GFP, 68.6% were mononucleate, 18.0% were binucleate and 13.4% had four nuclei. Likewise, in flies expressing RNAi#2, 44.3% of spermatids were mononucleate, 26.1% were binucleate and 29.6% had four nuclei whereas in flies co-expressing RNAi#2 and DE-cad-GFP, 67.5% were mononucleate, 20.3% were binucleate and 12.1% had four nuclei. Thus, expression of DE-cadherin partially bypasses the requirement for anillin during cytokinesis.

Figure 3.9. DE-cad-GFP suppresses cytokinesis defects caused by anillin depletion.

(A-D’) Fluorescence (A-D) and corresponding phase-contrast (A’-D’) micrographs of dividing spermatocytes expressing DE-cad-GFP. (A,A’) In wild-type cells DE-cad-GFP localizes to the cortex and in puncta at the poles prior to cytokinesis (early). As cytokinesis proceeds (mid), DE- cad-GFP begins to accumulate at the equator (arrows) (B,B’). During later stages of cytokinesis (late), DE-cad-GFP becomes highly concentrated in the furrow in wild-type (C,C’) and dsRNA- expressing cells (D,D’). Bar, 20µm. (E) Expression of DE-cad-GFP greatly reduces the percentage of multinucleate spermatids in flies expressing dsRNA directed against anillin but not in flies mutant for fwd. Suppression by DE-cad-GFP is observed for two independent β2t>anillin-RNAigenomic-cDNA fly lines (RNAi#1 and RNAi#2) and for UAS>anillin-RNAiVDRC (RNAi(VDRC)) flies. The number of spermatids counted for each genotype is indicated (n).

Figure 3.10. Membrane trafficking appears unaffected in anillin-depleted cells.

(A-D’) Fluorescence micrographs of dividing spermatocytes with corresponding phase contrast images (A’’-D’’). In wild-type cells, puncta containing clathrin-light-chain GFP (CLC-GFP) or a secreted GFP (sGFP) localize to the cleavage furrow (arrows) (A and C). This localization appears unaffected in dsRNA-expressing cells (B and D). (A’’-D’’) Corresponding merged images of CLC-GFP (green in A’’,B’’) or sGFP (green in C’’,D’’) with DNA (magenta). Bar, 20µm.

3.2.8 DE-cad-GFP recruits armadillo and α-catenin to the cleavage furrow

Because expression of DE-cad-GFP suppressed loss of anillin, I tested whether DE-cad and other members of the cadherin-catenin complex are normally expressed in developing male germ cells. Immunofluorescence experiments using specific antibodies revealed that β-catenin (armadillo), α-catenin and DE-cad are normally undetectable in dividing spermatocytes (Figure 3.11C, F and H; Figure 3.12A and B), whereas, in the same testis preparations, all three proteins were present in structures such as the apical hub of the testis and in epithelial cells of the seminal vesicle (Figure 3.11A, B, D, E and G). Strikingly, expression of DE-cad-GFP resulted in recruitment of armadillo and α-catenin to membrane structures including cleavage furrows of dividing cells (Figure 3.12C and D). In addition, Western blot analysis revealed increased armadillo and α-catenin protein levels in DE-cad-GFP expressing cells as compared to wild type (Figure 3.12E and F). Armadillo and α-catenin were not detected in spermatocytes expressing GFPNLS (not shown), demonstrating that catenin recruitment is not a secondary effect of GFP expression in male germ cells. Thus, DE-cad expression appears to stabilize and localize endogenous adherens junction proteins in dividing spermatocytes.

Figure 3.11. Endogenous DE-cadherin, armadillo and α-catenin are not detected in dividing spermatocytes.

(A-H’) Fluorescence micrographs of male reproductive structures stained for DE-cad (green in A’-C’), armadillo (Arm) (green in D’-F’), α-catenin (α-cat) (green in G’ and H’) and DNA (magenta). DE-cad, Arm and α-cat localize to membrane structures such as at the apical hub near the tip of the testis (arrows in A and D) and in the seminal vesicle (B,E,G) but are absent in dividing spermatocytes (C,F,H). Bar, 20µm.

Figure 3.12. DE-cad-GFP recruits armadillo and α-catenin to the cleavage furrow.

Fluorescence micrographs of dividing wild-type (A-A’,B-B’) or DE-cad-GFP-expressing (C- C’’,D-D’’) spermatocytes stained for armadillo (Arm) or α-catenin (α-cat). In wild type, Arm (A) and α-cat (B) are absent from the cleavage furrow (arrows). However, in cells expressing DE-cad-GFP (C,D), Arm (C’) and α-cat (D’) are clearly recruited to the furrow and to other regions of the plasma membrane. Arm also colocalizes with DE-cad in intracellular puncta. (A’,B’,C’’,D’’) Corresponding merged images showing DE-cad-GFP (green, C’’,D’’), Arm (red, A’,C’’), α-cat (red, B’,D’’) and DNA (blue). Bar, 20µm. (E,F) Immunoblots of whole testis proteins show increased levels of armadillo and α-catenin in DE-cad-GFP expressing male germ cells. DE-cad-GFP expressing cells have significantly higher levels of armadillo (E) (~105kDa) and α-catenin (F) (~110kDa) as compared to wild type. β-tubulin (~50kDa) was used as a loading control.

3.2.9 Adherens junction proteins can substitute for anillin in anchoring F- actin at the cleavage furrow

To determine if DE-cad-GFP expression suppresses loss of anillin by retaining F-actin at the cell equator, I examined F-actin localization in live cells using an mCherry fusion to the F-actin- binding calponin-homology domain of utrophin (mCherry-Utr-CH) (Burkel et al., 2007). During cleavage in wild-type and DE-cad-GFP-expressing cells, F-actin remained highly concentrated at the equator of the cell (Figure 3.13A and B, n=5/6 cells for wild type and n=4/5 cells for DE- cad-GFP). In contrast, in anillin-depleted cells, F-actin moved around the cell cortex, similar to the behavior of myosin II (compare Figure 3.13C to 3.7H). I observed different levels of phenotypic severity: cells were characterized as normal when localization of F-actin resembled controls, intermediate where there was some movement of F-actin from pole to pole, or severe when oscillations were dramatic and accompanied by the majority of F-actin leaving the area of the furrow. Of cells expressing dsRNA directed against anillin, none of the cells were normal, 2/7 were intermediate and 5/7 were severe (Figure 3.13C). In contrast, of cells depleted of anillin and expressing DE-cad-GFP, 1/7 were normal, 6/7 were intermediate (Figure 3.13D) and none were severe. Thus, like anillin, adherens junction proteins can restrict actomyosin contractility to the equator of the cell.

To determine if DE-cad-GFP substitutes for anillin by recruiting septins to the cleavage furrow, I examined peanut localization in anillin-depleted cells expressing DE-cad-GFP. Unlike wild-type cells, in which peanut was abundant at the cell equator (Figure 3.14A), cells expressing dsRNA directed against anillin had no detectable peanut at the equator regardless of whether DE-cad-GFP was co-expressed (Figure 3.14B and C). Thus, the data suggest that DE- cad-GFP promotes successful cytokinesis by providing an alternative means of anchoring F-actin and myosin at the cleavage furrow.

Figure 3.13. Anillin and DE-cadherin restrict F-actin to the cleavage furrow.

(A-D) Time-lapse images showing the change in localization of mCherry-Utr-CH (which binds F-actin) during spermatocyte cytokinesis. In control cells (A) and cells expressing DE-cad-GFP (B), mCherry-Utr-CH is tightly associated with the furrow, whereas in dsRNA-expressing cells mCherry-Utr-CH moves around the cell cortex in an oscillatory manner (C). DE-cad-GFP expression stabilizes mCherry-Utr-CH at the furrow in anillin-depleted cells (D). The images shown were taken at 40 sec intervals. Bar, 5µm.

Figure 3.14. DE-cadherin expression does not rescue peanut localization in anillin-depleted cells.

(A-C’) Fluorescence micrographs of dividing spermatocytes stained for peanut (Pnut) (red) and DNA (blue) with corresponding phase contrast images (A’’-C’’). Pnut localizes to the cleavage furrow (arrows) in wild-type cells (A) but not in dsRNA-expressing cells (B) or in cells expressing DE-cad-GFP (green in C’) and dsRNA (C). Bar, 20µm.

3.2.10 The ability of E-cadherin to suppress loss of anillin is conserved in mouse L-fibroblasts

To test whether classical cadherins are generally capable of suppressing loss of anillin function, our collaborators examined mouse L-fibroblast cells expressing E-cadherin under control of an inducible promoter (Angres et al., 1996; Goldbach et al., 2010). Knockdown of anillin with two different shRNAs caused a cytokinesis defect in cells grown in the absence of E-cadherin induction, resulting in 43 or 47% multinucleate cells (Figure 3.15). In contrast, expression of E- cadherin substantially suppressed the cytokinesis defect caused by anillin depletion, resulting in only 24 or 22% polyploidy. Thus, E-cadherin expression is sufficient to render mammalian cells less sensitive to loss of anillin.

Figure 3.15. Expression of E-cadherin suppresses cytokinesis defects caused by depletion of anillin in mouse L cells.

(A) Immunoblots showing reduced levels of anillin protein (~190 kDa) in anillin shRNA- expressing mouse LP (empty vector) or LE (E-cadherin expressing) cells (Anil KD) as compared to scrambled shRNA-expressing cells (Ctrl KD). GAPDH (~40 kDa) is the loading control. (B) Quantitation of multinucleate cells confirms suppression of anillin loss by expression of E- cadherin. *p value=0.00076. n is the number of experiments. 100 cells were counted in triplicate for each experiment. (C,D) Micrographs of LP and LE cells expressing scrambled shRNA (C,D) or anillin shRNA (C’,D’) marked by GFP (green). DNA is stained with Hoechst (blue). Representative multinucleate cells are marked (white arrowheads). Bar, 100µm. Modified slightly from figure assembled by Nolan Beise.

3.3 Conclusions

Anillin localizes to several actin-rich structures in the Drosophila testis and is required for male meiotic cytokinesis. Immunostaining with a newly generated anti-anillin antibody and expression of GFP- or RFP-anillin reveal anillin localization to cleavage furrows, ring canals, dense bodies and investment cones. Depleting anillin – either by RNAi or generation of mutant clones – causes defects in meiotic cytokinesis, as evidenced by the presence of multinucleate spermatids. In elongated spermatids depleted of anillin, investments cones are misshapen and myosin VI, which normally localizes to the leading edge of the cones, is diffuse or absent. This defect in individualization could be directly caused by the absence of anillin or it could be a secondary effect of earlier cytokinesis failures. Consistent with the latter idea, similar defects are observed in fwd mutant spermatids. However, an additional method of disrupting anillin function post-meiotically would be required to fully distinguish between these interpretations.

The results of experiments focused on meiotic cytokinesis support a model in which anillin acts to link the CR to the plasma membrane. FRAP analysis shows that anillin and septins are stably associated with the furrow. This is expected if anillin, possibly in conjunction with septins, acts as an anchor. Unexpectedly, I found that myosin II also does not undergo rapid exchange during constriction. This runs counter to current theories of CR dynamics that describe ring components as being highly dynamic (Section 1.1.1.2). Our results, along with a similar finding in C. elegans embryos (Carvalho et al., 2009), suggest a different model, at least for some metazoan cells, in which F-actin and myosin do not turn over but rather disassemble during furrow ingression.

Additional support for an anillin role in CR-membrane attachment comes from analysis of germ cells expressing dsRNA directed against anillin. Anillin-depleted spermatocytes exhibit late defects in cytokinesis – a cleavage furrow forms and ingresses, but in a high percentage of cells eventually regresses. Immunostaining or imaging of fluorescent markers reveals that septins are never recruited to the furrow, whereas F-actin and Rho are found in the furrow but are diffuse. FRAP analysis and live imaging of Sqh-GFP in anillin-depleted cells shows that myosin becomes highly dynamic and cycles around the periphery of the cell in the absence of anillin. A marker for F-actin shows similar dynamics in anillin-depleted cells, confirming that both major components of the actomyosin ring require anillin for proper localization during ingression.

Taken together, data presented here suggest that anillin, together with septins, is part of a stable complex at the furrow, where it is required to constrain F-actin and myosin II.

Anillin does not appear to play a role in membrane trafficking during cytokinesis. Localization patterns of three trafficking markers – sGFP, CLC-GFP and DE-cad-GFP – are indistinguishable between wild-type cells and cells expressing dsRNA directed against anillin. This suggests that, despite the late nature of the cytokinesis defect in anillin-depleted cells, anillin does not play a role in membrane trafficking. However, although trafficking itself does not appear to be impaired, it remains possible that anillin depletion results in defects in membrane fusion within the furrow that are not detected by the imaging methods described here.

DE-cadherin has the ability to suppress cytokinesis defects caused by loss of anillin. Expression of DE-cad-GFP significantly decreases the frequency of cytokinesis failure in spermatocytes expressing either of two dsRNAs directed against anillin. Although I was unable to detect endogenous DE-cadherin, armadillo or α-catenin in primary spermatocytes, when DE- cad-GFP is expressed ubiquitously, all three proteins are present in membrane structures including cleavage furrows. The F-actin marker mCherry-Utr-CH appeared more stable in furrows of cells co-expressing DE-cad-GFP and dsRNA directed against anillin, compared to cells expressing dsRNA alone. This suggests that the cadherin-catenin complex can partially replace anillin-septins in linking F-actin to the plasma membrane during ingression. This partial interchangeability of protein complexes appears to be evolutionarily conserved, as E-cadherin is able to suppress cytokinesis defects caused by anillin knockdown in mouse L-fibroblasts. The implications of these findings and proposed follow-up experiments are discussed further in the following chapter.

Chapter 4

Discussion and Future Directions

4 Discussion and Future Directions

In this chapter I discuss data presented in Chapter 3 and consider the implications for our current understanding of cytokinesis and developmental biology. Topics of discussion fall into three main sections: 1) the cytoskeleton-membrane crosslinking role of anillin during cytokinesis, 2) additional roles for anillin in Drosophila spermatogenesis and 3) potential contributions of E- cadherin to cytokinesis. At the end of each section I propose potential follow-up studies aimed at answering some of the questions raised therein. I begin by discussing the primary conclusion from the research presented in this thesis – that anillin acts to anchor the actomyosin ring to the plasma membrane during cleavage furrow ingression.

4.1 Anillin in contractile ring-membrane attachment

Successful cytokinesis requires anchoring of the contractile ring to the plasma membrane in the plane of division. Anillin has been postulated to play this role by recruiting septins and binding and stabilizing F-actin and myosin II during animal cell cytokinesis (reviewed in Eggert et al., 2006). Our results are consistent with this model, and show for the first time that anillin is required to restrict F-actin to the equator. Moreover, we find that anillin and the septins can be replaced by the cadherin–catenin complex to carry out this critical function. The interchangeability of these non-homologous cassettes suggests that two shared properties are required during cytokinesis: the ability to form a stable attachment to the plasma membrane and the ability to bind and bundle actin filaments. Let us begin with the first property.

4.1.1 Dynamics of contractile ring components during cytokinesis

In the prevailing model of CR dynamics, the major components, F-actin and non-muscle myosin II, turn over at a high rate during ingression. This is based on the results of several FRAP experiments performed in yeast and animal cells. Among the proteins shown to be highly dynamic in the furrow are myosin II heavy chain (Myo2p), essential light chain (Cdc4p) and regulatory light chain (Rlc1p), and the actin-binding proteins tropomyosin (Cdc8p) and IQGAP (Rng2p) in S. pombe (Pelham and Chang, 2002; Wong et al., 2002; Clifford et al., 2008); myosin II heavy chain (Myo1) in S. cerevisiae (Lister et al., 2006); and actin (tagged directly with GFP

80 or rhodamine) in mammalian cells (Guha et al., 2005; Murthy and Wadsworth, 2005).

My examination of myosin II dynamics in Drosophila spermatocytes, along with similar recent findings in the C. elegans embryo (Carvalho et al., 2009), provide evidence that contradicts this model of a highly dynamic actomyosin ring. In FRAP experiments presented in Section 3.2.4 I show that, in wild-type spermatocytes, Sqh-GFP fluorescence recovers at a much slower rate and to a lesser degree than PLCδ-PH-GFP. Carvalho et al. (2009) present similar data for myosin heavy chain (NMY-2) in dividing C. elegans embryos.

How do we resolve this apparent contradiction? The fact that similar results were obtained in Drosophila spermatocytes and C. elegans embryos suggests this is not an anomaly, but, rather, reflects a real and widespread aspect of actomyosin dynamics in animal cells. Should this be the case, it would represent a significant mechanistic difference between cytokinesis in animal cells versus budding and fission yeast. Whereas components of the CR turn over at a high rate in yeast, in at least some animal cells the CR does not turn over but rather is disassembled during constriction. Although there are many common features of cytokinesis between these divergent evolutionary branches, this may represent one of the distinctions (reviewed in Balasubramanian et al., 2004).

Do the results with fluorescently tagged actin mean that CR components are highly dynamic in mammalian tissue culture cells but not in Drosophila spermatocytes and C. elegans embryos? Probably not, since the mechanics of CR-driven cytokinesis are highly conserved in animal cells (Glotzer, 2001; Balasubramanian et al., 2004). Carvalho et al. (2009) suggest that because of the difficulty of incorporating tagged monomeric actin into formin-nucleated filaments, fluorescence in the furrow might actually represent short-lived Arp2/3-nucleated actin arrays; the majority of F-actin, existing in unbranched form, would thus be greatly underrepresented. I tend to agree with this interpretation. It would be interesting to test the dynamics of myosin in the same mammalian tissue culture cells used to probe F-actin dynamics (Guha et al., 2005; Murthy and Wadsworth, 2005). I anticipate that, similar to Drosophila and C. elegans, myosin would not exhibit rapid turnover during constriction. It is still possible that the actomyosin ring behaves differently in these cell types or that F-actin and myosin exhibit different dynamics from each other. Indeed there is some evidence for the latter. In a recent study, Zhou and Wang (2007) used total internal reflection fluorescence (TIRF) microscopy to

81 show that, at least for assembly of the actomyosin ring, F-actin and myosin exhibit markedly different spatial dynamics. Whereas myosin shows no directional flow during early cytokinesis, actin filaments undergo a striking flux toward the equator.

What about the dynamics of anillin and septins during furrow ingression? If these proteins act together to link the actomyosin ring to the plasma membrane (see below) we might expect them to associate tightly with the membrane. My results show for the first time that a metazoan anillin does not undergo rapid exchange during ingression (Section 3.2.4). Similar results were obtained for the anillin-like protein Mid1p in S. pombe (Clifford et al., 2008). Likewise, septins, including Sep2 in Drosophila (Section 3.2.4) and UNC-59 in C. elegans, do not exhibit rapid turnover (Carvalho et al., 2009). In S. cerevisiae, septins (four out of five septins were tested) are dynamic for a short temporal window during rearrangement of the septin ring, but become immobile again as cytokinesis proceeds (Dobbelaere et al., 2003). Interestingly, in S. pombe, the septin Spn4p does not exchange with cytoplasmic pools in wild- type cells but is highly dynamic in Δmid2 mutants (Berlin et al., 2003). This suggests that anillin-like proteins are required to keep septins attached to the membrane. Further evidence for this function of anillin is presented in the following section.

4.1.2 Anillin as a cytoskeletal-membrane crosslinker

Several lines of evidence support the conclusion that anillin links the contractile ring to the furrow membrane during constriction. Anillin appears to be at the centre of a molecular interaction network consisting of itself and four other central cytokinesis contributors: F-actin, myosin II, septins and phosphoinositides. I propose that anillin and septins act together to link F- actin and myosin II to phosphoinositides, and most likely PIP2, in the area of the furrow.

Anillin and septins might both contribute to plasma membrane binding through interactions with PIPs. Septins bind phosphoinositides in vitro and their localization is affected by alterations in phosphoinositide levels in vivo (Zhang et al., 1999; Casamayor and Snyder, 2003; Rodriguez-Escudero et al., 2005; Kouranti et al., 2006; Tanaka-Takiguchi et al., 2009) (Ho-Chun Wei, unpublished). Since plasma membrane-associated septin filaments form a gauze-like mesh that serves as a diffusion barrier (Takizawa et al., 2000; Schmidt and Nichols,

2004; Finger, 2005; Rodal et al., 2005), it is likely that binding of septins to PIP2-containing membranes in the cleavage furrow promotes formation of a stable molecular fence. Although

82 anillin has not been shown to bind phospholipids directly, the presence of the conserved PH domain (Oegema et al., 2000) suggests that anillin may also interact with membranes. Indeed, in septin knockdown cells, anillin still localizes to the membrane in a Rho-dependent manner (Straight et al., 2005; Hickson and O'Farrell, 2008; Piekny and Glotzer, 2008).

In addition to interactions with the membrane, anillin and septins might both contribute to interactions with the contractile ring. Anillin binds and bundles actin filaments and also binds myosin II, thereby interacting directly with both major contractile ring components (Field and Alberts, 1995; Straight et al., 2005). Recently, the mammalian septin, SEPT2, was also shown to interact directly with myosin II (Joo et al., 2007). We therefore propose that anillin stabilizes the cleavage furrow by recruiting septins and acting with septins to link the actomyosin ring to the membrane. Consistent with this, loss of anillin leads to destabilization of myosin II and F-actin, which move freely around the plasma membrane during cleavage of anillin-deficient cells (Sections 3.2.5 and 3.2.9) (Straight et al., 2005; Hickson and O'Farrell, 2008; Piekny and Glotzer, 2008). Indeed, our FRAP data confirm that myosin completely loses its stable association with the furrow in anillin depleted cells (Section 3.2.5). Anillin may also stabilize the contractile ring through indirect interactions with microtubule plus ends at the cell equator (D'Avino et al., 2008; Gregory et al., 2008; Hickson and O'Farrell, 2008). However, the fact that the cadherin-catenin complex can suppress loss of anillin in dividing spermatocytes suggests that the membrane-actin cross-linking properties of anillin may be sufficient to promote cytokinesis.

Suppression by E-cadherin also suggests that cells may exhibit differential sensitivity to loss of anillin in vivo, depending on the presence of alternative mechanisms to link F-actin and myosin to the plasma membrane. Other cytoskeletal-membrane crosslinkers, such as ERM proteins, localize to the cleavage furrow and could potentially carry out this function (Sato et al., 1991; Carreno et al., 2008; Kunda et al., 2008). In any case, it is likely that DE-cad-mediated suppression of the cytokinesis defects in anillin-depleted cells relies on a small amount of residual anillin and septins, since presumably these proteins are required for stabilization of post- division ring canals.

4.1.3 Proposal: Testing phospholipid-binding ability of anillin

In the model just presented, anillin and septins link the contractile ring to the plasma membrane. To test whether anillin has the ability to interact directly with the membrane, I propose a series of

83 experiments that focus on the PH domain. Although anillin fragments lacking the PH domain can localize to the cleavage furrow, the PH domain is required for robust localization and function (Oegema et al., 2000; Piekny and Glotzer, 2008). However, it remains unclear whether, in addition to its ability to recruit and bind septins, this domain can interact with PIPs (Oegema et al., 2000; Field et al., 2005a; D'Avino et al., 2008; Silverman-Gavrila et al., 2008). Indeed, there are no published reports describing the PIP-binding abilities of the anillin PH domain.

First, I propose examining the phospholipid-binding properties of the anillin PH domain in vitro using two complementary methods. The first is a phospholipid overlay assay, which involves passing recombinant protein over nitrocellulose membranes spotted with a range of immobilized lipids, including PI and its derivatives (PIP strips, Echelon Biosciences). This allows for detection of specific phospholipid-binding abilities in a semi-quantitative manner. The second method is a lipid flotation assay. This tests for the ability of recombinant protein to bind liposomes in solution and is thus considered more physiologically relevant (Kretzschmar et al., 1996; Skwarek et al., 2007). For lipid flotation assays, recombinant protein is mixed with liposomes containing a small percentage of the phospholipid being tested. Following ultracentrifugation in a sucrose density gradient, proteins that separate in liposome-containing fractions are detected by immunoblotting. A GST-fusion to a portion of anillin that contains the PH domain can be used for both methods. Although the PH domain is insoluble on its own (my unpublished results) a C-terminal fragment that includes the PH domain and a small amount of upstream sequence (GST-anillin-CT) is soluble and thus appropriate for these experiments (Silverman-Gavrila et al., 2008).

The functional importance of PIP-binding, should it be detected, can be examined by targeted mutation of positively charged residues in the anillin PH domain: lysine (K) or arginine (R) can be changed to glutamine (Q). Although these changes have little effect on protein folding, they can inhibit interactions with PIPs (Honing et al., 2005). I recommend mutating 1127-1128 RR to QQ. These arginine residues fall within the β2 strand of the anillin PH domain

(Field et al., 2005) and positive residues in the β1/β2 loops of diverse PH domains have been shown to be required for interactions with PIPs (reviewed in Rebecchi and Scarlata, 1998; Lemmon et al., 2002). R1128 specifically is likely to be important for PIP interactions since, in sequence alignments, it aligns with a key arginine residue found in the β2 strand of PLC-δ1, Btk and PDK1 PH domains (Ferguson et al., 1995; Mattsson et al., 1996; Komander et al., 2004).

Furthermore, two of the point mutations identified in strong maternal effect alleles of Drosophila anillin fall in β1/β2 loop of the PH domain (Field et al., 2005). The fact that the mutated anillin protein encoded by these alleles shows less robust localization to furrow canals in cellularizing embryos suggests that the β1/β2 loop may be important for interaction with PIPs in the furrow 1127-1128 1147-1149 membrane. Although not as strong candidates as RR , KRK in the β3/β4 loop and 1175 R in the β5/β6 loop may also contribute to PIP interactions and could also be tested.

The ability of the mutated constructs to interact with PIPs can be tested in vitro using the methods discussed above. If PIP-binding is abolished, a full length RFP-anillin construct can be generated containing the relevant point mutations (RFP-anillinmutPH). The construct would be based on RFP-anillin that rescues anillin mutant germline clones (Section 3.2.2). RFP- mutPH anillin can be expressed in S2 cells, or in male germ cells using the β2-tubulin promoter. The importance of PIP-binding for anillin localization can be examined in both systems. In addition, RFP-anillinmutPH can be tested for its ability to rescue the anillin cytokinesis phenotype in male germ cells. If PIP-binding properties of anillin are shown to be important for function, this would contribute greatly to a model in which anillin anchors the contractile ring to the furrow membrane.

4.2 Anillin function(s) in Drosophila spermatogenesis

Anillin may have additional roles beyond cytokinesis in vivo. However, it is difficult to obtain cells entirely lacking anillin protein, and earlier defects in cytokinesis may obscure later phenotypes. For example, although we observed defects in actin cone formation in anillin- depleted spermatids (Section 3.2.3), this could reflect either a specific role for anillin in individualization or an indirect effect of cytokinesis failure on membrane topology. Similarly, persistent association of anillin with ring canals (Section 3.2.1) suggests a role in ring canal stability. However, since ring canals are formed as a by-product of contractile rings, disrupting cytokinesis would prevent their formation. One method of studying anillin function at later stages of male germ cell development could involve using temperature-sensitive alleles. Instead of the more laborious technique of identifying temperature-sensitive alleles through genetic screening, a temperature-sensitive version of anillin could be generated by insertion of a temperature-dependent, self-splicing intein (Zeidler et al., 2004). Inteins are peptide equivalents of introns in genes. They can be inserted into functional regions of proteins and their removal,

85 which occurs through a self-splicing mechanism, results in regeneration of normal protein structure and function. In this case, flies can be generated expressing an intein-containing anillin construct as their only copy of anillin. At various times after transferring flies from the permissive to restrictive temperature, male germ cells can be dissected out and examined for potential post-meiotic defects. Alternatively, one could employ other techniques to destabilize the anillin protein dynamically in living cells (see Section 4.2.4).

4.2.1 Individualization

Anillin may have a role in Drosophila spermatid individualization. Anillin localizes to the leading edge of actin-based investment cones in a manner similar to the unconventional myosin VI (Section 3.2.1). In male germ cells expressing dsRNA directed against anillin, investment cones form but are misshapen (Section 3.2.3), suggesting a role for anillin in investment cone structural stability. However, the fact that similar defects are observed in fwd spermatids indicates that this may be a secondary defect of earlier failures in cytokinesis. Specifically, the increased size of multinucleate spermatids could be a source of steric hindrance to investment cones attempting to translocate caudally. Alternatively, fwd and anillin might both contribute to proper investment cone function. Hence, a genuine role for anillin in individualization could be obscured by prior cytokinesis failure. In order to distinguish among these possibilities, anillin function must be disrupted after the meiotic phase in development.

If anillin does contribute to investment cone function, what role might it play? All anillin functions described to date are in contractile events involving F-actin and non-muscle myosin II (Section 1.2.3). However, the contribution of anillin to these processes does not appear to be in contraction per se – that is, in the mechanical motions of myosin acting on antiparallel actin filaments. Instead, the results of this and other studies suggest anillin acts as a multidomain scaffolding protein, linking F-actin to other structural and regulatory molecules. Thus, anillin could bind and bundle F-actin in investment cones and/or act as a link to myosin VI or plasma membrane PIPs.

4.2.2 Anillin and septins in ring canal stability

In addition to individualization, anillin may play a functional role in ring canals. Anillin localizes to mitotic and meiotic ring canals and maintains its association throughout spermatid

86 elongation (Section 3.2.1) (Hime et al., 1996). Importantly, its localization coincides with septins – peanut, Sep1 and Sep2 (Hime et al., 1996). Septin filaments are sometimes described as cytoskeletal-like (Ihara et al., 2005; Caudron and Barral, 2009) and may be acting as a type of cytoskeleton in this context.

Septins have the intrinsic ability to form into rings as observed in vitro. A mammalian septin complex of SEPT2/6/7 can self-assemble into rings with a diameter of ~0.6 µm in the absence of actin (Kinoshita et al., 2002) and a recombinant yeast complex of Cdc3/11/12/10 self- assembles into rings with a diameter of ~1 µm (Farkasovsky et al., 2005). Kinoshita et al. (2002) suggest that in vitro rings of septin filaments resemble similar in vivo septin formations in cellular structures such as intercellular bridges that form after cytokinesis in cultured mammalian cells (diameter ~0.5 µm; Mullins and Biesele, 1977) and Drosophila male ring canals (1–1.5 µm; Hime et al. 1996). Other examples include the bud neck in S. cerevisiae (diameter, 0.42–0.54 µm; Byers and Goetsch, 1976) and the stalks that connect Drosophila blastomeres to the yolk sac after cellularization (~0.5 µm; Robinson and Cooley, 1996). Thus, energetically favourable septin rings could play a cytoskeletal-like role in maintaining intercellular bridges such as ring canals.

I therefore propose that anillin plays a structural role in maintaining septin-based ring canals. It might accomplish this by cooperating with septins in membrane binding (see above) or by binding additional structural or regulatory ring canal components. In addition, anillin might be required to bind F-actin that associates with ring canals in fusomes or in elongating sperm tails. As is the case for studying anillin function in individualization, disruption of anillin post- meiotically will be required to establish such functions.

4.2.3 Anillin stability and regulation

My investigations into anillin function in the testis suggest that anillin protein may be unstable in developing male germ cells. One of the RNAi approaches used to knock down anillin in the testis was expression of UAS>anillin-RNAiVDRC under control of the germline specific Bam- Gal4 driver. This resulted in a high frequency of meiotic cytokinesis failure (Section 3.2.2). Unexpectedly, Bam-Gal4-driven expression of a large number of dsRNAs directed against other genes did not result in any detectable phenotypes. This included dsRNAs directed against fwd,

87 sktl, all five Drosophila septins and several additional PIP-regulating genes (Lacramioara Fabian and my unpublished results).

There are a number of possible explanations as to why Bam-Gal4-driven dsRNA directed against anillin caused detectable defects while other dsRNAs did not. Perhaps the most likely possibility is that anillin is less stable in developing male germ cells than other proteins tested. If some or all of the protein is turned over during spermatocyte growth, then dsRNA-mediated knockdown would have a stronger effect. Alternatively, protein levels or stoichiometry might matter more for anillin than for other proteins. A perhaps less likely possibility is that anillin dsRNA might be expressed at a higher level or be more stable than other dsRNAs, resulting in more effective knockdown.

If anillin does turn over at a high rate, this result may suggest something interesting about anillin regulation. Does all of the anillin in the cell turn over at the same rate, or is a subset more stable – for example, nuclear anillin versus anillin in ring canals? Anillin stability could be regulated by secondary modifications such as phosphorylation and ubiquitination. There is some evidence that Xenopus anillin is phosphorylated and APC/C-mediated ubiquitination of human anillin is required for degradation after cell division (Straight et al., 2005; Zhao and Fang, 2005). For more on anillin regulation in Drosophila male germ cells, including data demonstrating that anillin is phosphorylated, see the Appendix.

4.2.4 Proposal: Examining post-meiotic functions of anillin

Studying the function of anillin in post-meiotic male germ cells presents a number of difficulties. Any method of targeted anillin disruption that relies on dsRNA or protein being synthesized pre- meiotically – for example, RNAi-mediated knockdown approaches described in this thesis – has a high probability of causing cytokinesis defects. Later developmental defects may thus be obscured.

One way to get around this would be to knock down anillin function using a transgene whose product is translated post-meiotically. Although almost no transcription occurs post- meiotically, there is a family of genes whose translation is inhibited until after meiosis (Kuhn et al., 1988; Schafer et al., 1990; Yanicostas and Lepesant, 1990; Fuller, 1993). Translational timing is controlled by a conserved element of twelve nucleotides in the 5’ UTR called a

88 translational control element (TCE) (Schafer et al., 1990). Deletion of the TCE in Mst87F, a member of this gene family, causes a shift in translation from post- to pre-meiotic.

To specifically disrupt anillin function post-meiotically, I propose using a TCE in conjunction with in vivo tobacco etch virus (TEV) protease-mediated cleavage. This technique makes use of a seven-amino-acid site – ENLYFQS – recognized by TEV protease, which can be inserted into a protein of interest (Carrington and Dougherty, 1988; Dougherty et al., 1989). TEV protease-mediated cleavage has recently been validated as an effective means of spatially and temporally targeting protein cleavage in Drosophila (Harder et al., 2008; Pauli et al., 2008).

Two working parts are required for this method: 1) the TEV protease transgene containing a 5’ UTR TCE and 2) a full-length anillin construct containing internal TEV cleavage sites. Using standard molecular techniques, the 5’ UTR of Mst87F can be fused with the coding region of TEV protease and inserted into a vector containing the β2-tubulin promoter. In conjunction with this, a DNA sequence encoding the TEV site can be engineered into RFP- anillin, fused to the endogenous anillin promoter and inserted into the standard Drosophila transformation vector pCaSpeR4. In addition, a FLAG epitope tag can be fused to the C- terminus so that stability of N- and C-terminal fragments can be monitored. The TEV site should be inserted in a region of low sequence conservation, for example, between the anillin homology region and upstream sequence, to ensure proper protein folding. Several different TEV insertions should be designed and tested for efficacy of cleavage and for rescue of anillin mutant flies.

The RFP-anillin-FLAG constructs with cleavage sites can first be tested for effective cleavage in S2 cells expressing TEV protease (Schafer et al., 1990). Should cleavage occur, I expect the N-terminal fragment would no longer localize to the cortex in metaphase and only weakly to cleavage furrows, in a manner similar to a GFP-anillin construct lacking the PH domain in mammalian cells (Oegema et al., 2000). It is also possible that one or both protein fragments could be degraded following cleavage. In addition to determining localization in cells, successful cleavage and protein stability can be tested by immunoblotting with anti-anillin and anti-FLAG antibodies.

Vectors containing TEV protease and cleavable RFP-anillin-FLAG can then be used to generate transgenic fly lines. Cleavable RFP-anillin-FLAG should be tested for ability to rescue strong anillin alleles such as anilPZ and anil8. RFP-anillin-FLAG without cleavage sites should also be tested to ensure the C-terminal FLAG-tag does not interfere with anillin function. Male flies can then be generated that express cleavable RFP-anillin-FLAG as their only source of anillin, in combination with post-meiotically expressed TEV protease. If cleavable RFP-anillin- FLAG cannot rescue anillin alleles, the experiment can still be performed in male germline mutant clones.

Effects of disrupting anillin post-meiotically can then be examined. First, flies should be tested for fertility. If fertility is affected, male germ cells can then be examined morphologically and by immunostaining for defects in processes such as spermatid elongation and individualization. A positive result would extend knowledge of anillin function beyond cytokinesis and cytokinesis-like processes. For example, since anillin and septins are highly expressed in both diseased and normal CNS tissues (Hall et al., 2005a; Hall et al., 2005b), future studies using such techniques may uncover exciting new roles for anillin and the septins in the CNS.

4.3 A role for E-cadherin in animal cell cytokinesis?

Ectopic expression of E-cadherin can suppress loss of anillin in dividing cells. This unanticipated result raises several interesting questions. For example, what is the molecular mechanism of suppression by E-cadherin and does E-cadherin normally contribute to animal cell cytokinesis? Although a few studies report an examination of E-cadherin in cells undergoing cytokinesis, a direct functional contribution has yet to be described. In this section I discuss the link between E-cadherin and cytokinesis, beginning with an exploration of molecular mechanism.

4.3.1 Mechanism of E-cadherin suppression

Expression of DE-cadherin partially restores cytokinesis to anillin-depleted spermatocytes (Section 3.2.7), suggesting that the cadherin-catenin complex, like the anillin-septin cassette, is capable of linking the actomyosin ring to the furrow membrane. Classical cadherins such as DE- cadherin are transmembrane proteins that form stable membrane attachments by virtue of their

90 ability to mediate cell-cell adhesion (Takeichi, 1995). Ectopically expressed fluorescent E- cadherin and DE-cadherin fusion proteins localize to the cleavage furrow, where they associate with β-catenin (armadillo) and α-catenin during cytokinesis (Bauer et al., 2008) (Section 3.2.8). Homodimers of α-catenin bind and bundle actin filaments (Weis and Nelson, 2006; Hartsock and Nelson, 2008), suggesting that α-catenin is ideally suited to substitute for anillin during cytokinesis. Importantly, our collaborators found that expression of E-cadherin in mouse L-cells renders them less sensitive to loss of anillin (Section 3.2.10), supporting this idea. Indeed, our finding that DE-cadherin and E-cadherin behave similarly in this respect suggests that cells expressing adherens junction proteins may be less sensitive to loss of anillin during animal development.

Our data show that classical cadherins can stabilize F-actin in the contractile ring (Section 3.2.9). It is also possible that the cadherin-catenin complex may be stabilizing a connection between the central spindle and the furrow membrane through its ability to associate with microtubules (reviewed in Harris and Tepass, 2010). Recent studies suggest that adherens junctions have the ability to stabilize microtubule plus ends (Waterman-Storer et al., 2000; Ligon et al., 2001) as well as minus ends (Chausovsky et al., 2000; Meng et al., 2008). Moreover, adherens junctions are implicated in positioning the spindle during the asymmetric divisions of Drosophila sensory organ precursor cells and male germline stem cells (Le Borgne et al., 2002; Yamashita et al., 2003). It remains to be seen whether cadherin-catenins can stabilize microtubule plus ends in the vicinity of the furrow during conventional cytokinesis.

To summarize, our data suggest that, in the absence of anillin and septins, the cadherin- catenin complex can interact with and stabilize F-actin in the cleavage furrow (Figure 4.1B). However, it is also possible that cadherins stabilize apposed membranes between daughter cells through homophilic interactions, and that F-actin stabilization is a secondary effect (Figure 4.1C). These scenarios are not mutually exclusive and both may contribute (Figure 4.1D). A method of distinguishing between the possibilities is discussed in Section 4.3.3.

Figure 4.1. Cadherin-catenins can replace anillin-septins in stabilizing the furrow during cytokinesis.

(A) Schematic diagram of an animal cell undergoing cytokinesis with part of the furrow region (boxed area) magnified in panels B-E. (B) Anillin and septins (green) are required to anchor the actomyosin ring (red) to the furrow membrane during constriction. (C-E) The cadherin-catenin complex (blue) can suppress loss of anillin and septins by binding F-actin (C), stabilizing apposed membranes through cadherin homophilic interactions (D) or both (E).

4.3.2 An endogenous role for E-cadherin in cytokinesis?

Our experiments raise the question of whether cadherin-catenin function is normally important for cytokinesis. The fact that I was unable to detect endogenous DE-cadherin, armadillo or α-catenin in wild-type spermatocytes suggests these proteins do not contribute to cytokinesis in these cells. It is still possible that small amounts of protein are present but below detectable levels. Possible functions for endogenous cadherin-catenins could be probed by generating male germline clones mutant for armadillo or α-catenin using the method described above (see Section 2.3).

Outside of the spermatocyte context, only a few studies suggest a link between cadherins and cell cleavage. In the early zebrafish embryo, daughter blastomeres transition from being

92 loosely associated to having tightly apposed membranes. This is accomplished following ingression via delivery of adhesion proteins including E-cadherin and β-catenin to newly formed daughter membranes (Jesuthasan, 1998; Li et al., 2006). Disruption of cadherin-mediated adhesion causes defects in blastomere cohesion, but not in cytokinesis (Jesuthasan, 1998). In mouse fibroblasts, the introduction of E-cadherin-α-catenin fusion proteins has the opposite effect, causing a slight inhibition of cytokinesis (Nagafuchi et al., 1994). These fusion proteins likely form unregulated junctions that prevent cells from rounding up during division, suggesting that cells must coordinate the disassembly of adherens junctions with cytokinesis. Hence, the cadherin-catenin complex may not be required for cytokinesis per se, but is involved in changes in tissue architecture that must be tightly linked with cytokinesis.

Bauer et al. (2008) showed that ectopically expressed E-cadherin-Venus fusion proteins accumulate in cleavage furrows of cultured MDCK cells. They used E-cadherin-Venus as a representative of transmembrane proteins in general and did not ascribe to it a specific function. It is interesting to note the difference between the localization of E-cadherin fusion proteins described by Bauer et al. in MDCK cells with what we observe in Drosophila spermatocytes. Whereas E-cad-GFP localizes to large internal puncta and plasma membrane in spermatocytes (Figure 3.9A-C), E-cadherin-Venus is restricted to the plasma membrane in MDCK cells. This may reflect real differences in the role of membrane trafficking in different cell types. In dividing spermatocytes, E-cad-GFP may be trafficked from recycling endosomes to the plasma membrane under control of the exocyst complex, as has been shown in Drosophila epithelial cells (Langevin et al., 2005), and, or in addition, may travel through the secretory pathway.

Thus, further studies are required to determine whether classical cadherins normally contribute to animal cell cytokinesis. Our experiments in Drosophila spermatocytes and mouse L-fibroblasts strongly suggest they have this capability. In previous studies of cadherin function, subtle effects on cytokinesis may have been obscured by more obvious defects in tissue architecture. Therefore, precise methods of disrupting cadherin-catenin complexes in time and space will likely be required to separate specific cytokinesis-related functions from general roles in cell adhesion.

4.3.3 Proposal: Determining the mechanism of E-cadherin suppression

To distinguish among different models of anillin suppression by E-cadherin (Figure 4.1), I propose disrupting specific functional regions of DE-cadherin. I would generate flies that express one of two constructs: one that abolishes homophilic interactions in the extracellular region without affecting catenin binding and one that abolishes catenin binding without affect homophilic interactions. Such constructs exist and have already been validated.

A construct termed DE-cad-dCR3h (dCR = deletion of cadherin repeats) contains an internal deletion in the extracellular region of DE-cadherin (Oda and Tsukita, 1999). When expressed in S2 cells, DE-cad-dCR3h localizes to the plasma membrane but is unable to mediate cell aggregation. Expression of DE-cad-dCR3h-GFP in shg mutant flies results in only partial rescue of defective developmental processes such as tracheal morphogenesis. Importantly, DE- cad-dCR3h-GFP is found in aggregates with α-catenin, demonstrating that it retains its ability to associate with catenins.

Another construct, termed DE-cad-Δβ, contains a deletion of the C-terminal armadillo interaction domain (Pacquelet et al., 2003). When expressed in S2 cells, DE-cad-Δβ localizes to the plasma membrane and can still promote cell aggregation. Expression of DE-cad-Δβ in shg mutant flies results in little or no rescue of several oogenesis and embryogenesis related defects. Importantly, defects seen in flies expressing DE-cad-Δβ as their only copy of DE-cadherin resemble those seen in armadillo mutants (Peifer, 1993; Pacquelet et al., 2003), suggesting that the C-terminal DE-cadherin deletion specifically affects cadherin-catenin complex assembly.

In order to compare the effects of expressing different DE-cadherin constructs in male germ cells, all constructs should be generated in the same transformation vector. Since mutated forms of DE-cadherin have the potential to act as dominant negatives when expressed ubiquitously, I recommend limiting expression to male germ cells using a β2-tubulin vector. Full-length DE-cad-GFP (Oda and Tsukita, 2001) should be subcloned into this vector, as well as DE-cad-dCR3h-GFP and DE-cad-Δβ. In addition, a C-terminal GFP tag would need to be fused to DE-cad-Δβ (DE-cad-Δβ-GFP).

This experiment could also be done in L-fibroblasts. Similar constructs have been described for mammalian E-cadherin: one lacks the β-catenin binding region just as in flies and,

94 in the other, the extracellular cadherin repeats are replaced with the extracellular region of interleukin-2 receptor α subunit (IL2R) (Gottardi et al., 2001). Both constructs should be subcloned into the same inducible vector used for full-length E-cadherin expression (Angres et al., 1996; Goldbach et al., 2010).

Whether in Drosophila male germ cells or in mouse L-fibroblasts, the different cadherin constructs could be tested, as described, for their ability to suppress cytokinesis defects caused by expression of dsRNA directed against anillin. There are a number of different potential outcomes in these experiments. Taking the Drosophila constructs as examples, if DE-cad- dCR3h-GFP suppresses defects caused by dsRNA directed against anillin but DE-cad-Δβ-GFP does not, this would suggest that cadherin-catenins contribute by binding F-actin (Figure 4.1B). Conversely, if DE-cad-Δβ-GFP suppresses but DE-cad-dCR3h-GFP does not, this would suggest that extracellular homophilic interactions between cadherins are responsible for suppression (Figure 4.1C). If both homophilic interactions and F-actin-binding are involved (Figure 4.1D), one might expect to see lower levels of suppression, or none at all, by both constructs. Overall, if, as discussed in Section 4.3.2, classical cadherins normally contribute to successful animal cell cytokinesis, these experiments have the potential to explain how this function is carried out.

4.4 Concluding remarks

In summary, data presented here further establish anillin as a central organizing factor in cytokinesis. They also hint at additional developmental roles for the protein. Future experiments, such as those described in this chapter, will be required to determine if anillin does indeed have non-cytokinesis functions in development.

The finding that expression of E-cadherin can suppress cytokinesis defects in anillin- depleted cells nicely illustrates the potential for interchangeability of protein complexes. In this case, it appears that either anillin-septins or cadherin-catenins can anchor the contractile ring to the membrane during cleavage furrow ingression. This is a clear example of built-in redundancies providing a source of robustness and adaptability to cells. It also highlights the fact that evolution may take advantage of proteins or protein complexes with useful physical properties and apply them in new or slightly different contexts.

This is an exciting time for the cytokinesis field. The last decade or two has seen an explosion of data resulting in increasingly more mechanistic molecular models. Over the coming years, a combination of large scale screens and focused studies will undoubtedly fill in many of the gaps. In addition, more powerful computational methods will be of great help in testing theoretical models. Aside from providing great insights into this universal aspect of cell biology, these studies hold the potential to lead to targeted therapies in treating diseases such as cancer.

Appendix

Anillin Regulation and Phosphorylation in Drosophila Male Germ Cells

Abstract

In Chapter 3 of this thesis I presented the results of experiments focused on anillin function in Drosophila male germ cells. Here, I focus on the regulation of anillin in this system. I begin by describing a link between anillin and the cell cycle inhibitory kinase dMyt1. In myt1 mutants, anillin is aberrantly localized to the cell cortex in early spermatids. In addition, immunoblotting experiments reveal a change in the electrophoretic mobility of anillin protein species from myt1 testis lysates. The anillin species of higher apparent molecular weight, which is reduced in abundance in myt1 mutant testes, represents a phosphorylated form of the protein, as demonstrated by an in vitro phosphatase assay. Moreover, colocalization of anillin and phosphotyrosine epitopes in multiple spermatogenesis-related structures raises the possibility that anillin is developmentally regulated by phosphorylation of tyrosine residues.

Introduction

Anillin localization is tightly regulated in time and space during the cell cycle. Anillin is sequestered in the nucleus during interphase; during metaphase it relocalizes to the cortex; and shortly thereafter it becomes focused in the cleavage furrow (Section 1.2.2). Anillin regulation is likely controlled, at least in part, by post-translational modifications such as phosphorylation and ubiquitination. Indeed, ubiquitination by APC/C results in degradation of mammalian anillin at the end of cell division (Zhao and Fang, 2005).

In S. pombe, phosphorylation of the anillin-like protein Mid1p appears to play an integral role in its regulation. Like Drosophila and vertebrate anillins, Mid1p is nuclear during interphase. Its relocalization from the nucleus to the cell cortex is concurrent with protein hyperphosphorylation (Sohrmann et al., 1996). Several lines of evidence suggest that Mid1p is phosphorylated by the polo-like kinase Plo1p. mid1 and plo1 mutants exhibit similar defects in medial ring placement and mid1 plo1 double mutants resemble the single mutants (Bahler et al., 1998), suggesting they act in the same pathway. In addition, two-hybrid assay results suggest that Mid1p and Plo1p physically interact. Moreover, in plo1 mutants, Mid1p is unable to exit the nucleus (Bahler et al., 1998; Paoletti and Chang, 2000). Interestingly, Plo1p appears to regulate the cytoplasmic function of Mid1p as well as its nuclear exit, as a nuclear localization-defective

98 version of Mid1p localizes to the medial ring but is unable to rescue plo1 mutants (Paoletti and Chang, 2000). Despite the intimate connection between Mid1p and Plo1p, direct phosphorylation of Mid1p by Plo1p has yet to be demonstrated.

There is some evidence that Drosophila and vertebrate anillins are also regulated by phosphorylation. In the initial characterization of Drosophila anillin, the existence of a closely spaced doublet recognized by anti-anillin antibody in immunoblotting experiments suggested the possibility of post-translational modifications (Field and Alberts, 1995). The anillin species with slightly lower mobility is more prominent in extracts from early embryos – a developmental period coincident with high levels of cell division. In addition, Straight et al. (2004) cite unpublished data demonstrating that Xenopus anillin is phosphorylated by PLK1.

Polo-like kinases are among several kinases involved in regulating entry into mitosis or meiosis. Another important regulator is the inhibitory kinase Myt1, which acts on cyclin- dependent kinase 1 (Cdk1). A complex of Cdk1 and cyclin B provides a key signal to the cell to enter mitosis. Myt1, and the related Wee1 kinase, maintain Cdk1 in an inactive state to prevent premature entry. In S. pombe, Drosophila, vertebrates, echinoderms and plants, Cdk1 is phosphorylated on two inhibitory residues – Y15 and T14 (reviewed in O'Farrell, 2001; Lindqvist et al., 2009). In metazoans, Wee1 phosphorylates Y15 with high specificity (Parker et al., 1992) while Myt1 can phosphorylate both Y15 and T14 (Mueller et al., 1995). Whereas Wee1 is nuclear, Myt1 is cytoplasmic and membrane-associated. Myt1 and/or Wee1 keep Cdk1 inactive until Cdc25 phosphatase removes the phosphates from the inhibitory sites. Cdk1 then provides positive feedback by phosphorylating Cdc25 and Wee1/Myt1, increasing activation and inhibition respectively.

Drosophila has only one representative of each kinase, dMyt1 and dWee1. dWee1 is required for controlling mitotic entry during the rapid S/M nuclear cycles of early embryogenesis (Price et al., 2000; Stumpff et al., 2004). dMyt1 and dWee1 appear to be partially redundant in a number of contexts, although dMyt1 is the major Cdk1 inhibitory kinase in wing imaginal disc development and gametogenesis (Price et al., 2002; Jin et al., 2005; Jin et al., 2008). During spermatogenesis, dMyt1 regulates multiple aspects of cell cycle behaviour (Jin et al., 2005). In myt1 mutants, male germ cells inappropriately enter S- and M-phase, as evidenced by increased BrdU incorporation and phospho-histone H3 staining respectively. Germline-associated cyst

99 cells also undergo ectopic division. In addition to inappropriate timing of division, male germ cells occasionally undergo an extra round of mitotic division resulting in cysts of 128 spermatids instead of the usual 64. myt1 mutant phenotypes are phenocopied by expression of heat shock- inducible, non-inhibitable Cdk1, confirming that dMyt1 mainly acts through inhibition of Cdk1 activity. Thus, dMyt1 inhibition of Cdk1 is required for efficient coupling of cell differentiation with cell cycle progression during spermatogenesis.

Although phosphorylation by major serine/threonine kinases such as Cdk1, Aurora B and Polo is crucial during cell cycle regulation, phosphorylation on tyrosine residues also appears to be important during Drosophila gametogenesis. Tyrosine phosphorylated protein(s) localize to ring canals in male and female germ cells. In the ovary, ring canals become lined with phosphotyrosine epitopes as cytokinesis halts (Robinson et al., 1988). Two tyrosine kinases, Src64 and Tec29, are required for growth of actin-rich ring canals (Cooley, 1998; Dodson et al., 1998; Guarnieri et al., 1998). Tec29 localization and function are mediated by Src64 phosphorylation and PIP3 binding (Lu et al., 2004). In the testis, male germline ring canals also contain phosphotyrosine epitopes (Hime et al., 1996). In dividing spermatocytes, phosphotyrosine-containing proteins first appear as punctate dots along the contractile ring, later coalescing into a continuous ring as constriction proceeds.

In addition to functioning in ring canals, there is some evidence that tyrosine phosphorylation is important for cytokinesis itself. Src tyrosine kinase localizes to the cleavage furrow in mammalian cells and sea urchin embryos (Ng et al., 2005; Kasahara et al., 2007; Shafikhani et al., 2008). In HeLa cells, Src activity is required for ERK/MAPK activation at the midbody and successful abscission (Kasahara et al., 2007). In sea urchin embryos, Src is associated with detergent resistant membranes (DRMs), sometimes referred to as lipid rafts (Ng et al., 2005). Notably, DRMs isolated from dividing cells contain significantly higher levels of tyrosine phosphorylated Src and PLCδ than DRMs from interphase cells. In addition to their described role in female ring canal formation, the Drosophila tyrosine kinases Src64 and Tec29 play a role in contractile events during cellularization (Thomas and Wieschaus, 2004). In src64 and tec29 mutant embryos, the depth of membrane invagination is non-uniform and basal closure of the cells does not occur. It remains to be seen whether these kinases play a role in conventional Drosophila cytokinesis.

100

Materials and Methods

Unless otherwise indicated, experiments were performed as described in Chapter 2.

Drosophila genetics Flies used in myt1 experiments were myt11/Df(3L)64D-F hemizygotes (Garcia-Bellido et al., 1994; Jin et al., 2005).

Fluorescence microscopy

Testis dissections and immunostaining were performed as described (Section 2.4). To detect phosphotyrosine-containing epitopes, mouse anti-pTyr (4G10, Millipore) was used at 1:100.

Phosphatase assay and immunoprecipitation

Thirty pairs of wild-type testes were dissected in testis isolation buffer (TIB) (Casal et al., 1990) containing protease inhibitors (Complete Protease Inhibitor Tablets, Roche). Testes were transferred to 0.1 ml glass homogenizer tubes (Jencons Scientific) in 30 µl TIB with 1% Triton X-100 and protease inhibitors and homogenized manually. Homogenized materials were transferred to an eppendorf tube, centrifuged and supernatant was removed. In earlier experiments, anillin was determined to be insoluble in lysates containing detergent. Pellets were resuspended in 30 µl lambda phosphatase buffer with or without 400 units of lambda protein phosphatase (NEB). Reactions were carried out at 30˚C for 60 minutes. Samples were then boiled for 5 minutes in SDS-PAGE sample buffer, spun down and loaded onto an 8% SDS polyacrylamide gel. Immunoblotting was performed as described (Section 2.6).

Results

Anillin is aberrantly localized in male germ cells from myt1 mutant flies

As part of a collaboration to discover the function of dMyt1 in male germ cell development (Jin et al., 2005), we obtained preliminary results suggesting that the distribution of anillin and F- actin was abnormal in myt1 mutant males (J. Brill, unpublished observations). To examine this

101 further, I performed immunostaining experiments on male germ cells from wild-type and myt1/Df flies. Anillin and peanut localization appeared normal in pre-meiotic germ cells (not shown) and in dividing spermatocytes, where they colocalized at the cleavage furrow (Figure A1.1A and B). However, in early spermatids, localization patterns often appeared dramatically altered. Whereas, in wild type, anillin and peanut colocalized in ring canals (Figure A1.1C), in myt1/Df spermatids, anillin was found predominantly on the cortex and peanut was largely cytoplasmic (Figure A1.1D). In addition, F-actin, normally found in the fusome (Figure A1.1E), colocalized with cortical anillin in myt1/Df spermatids (Figure A1.1F). Thus dMyt1 is required for proper localization of anillin, peanut and F-actin in early spermatids.

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Figure A1.1. Anillin is mislocalized in myt1 mutant male germ cells.

(A-D) Male germ cells stained for anillin (red), peanut (green) and DNA (blue). (A,B) In dividing wild-type (A-A’’’) and myt1/Df (B-B’’’) spermatocytes, anillin (A,B) and peanut (A’,B’) colocalize (A’’,B’’) in cleavage furrows. Furrows are visible in corresponding phase-

103 contrast micrographs (A’’’,B’’’). (C) In early wild-type spermatids, anillin (C) and peanut (C’) colocalize (C’’) in ring canals (arrowhead). (D) Strikingly, in myt1/Df early spermatids, anillin (D) is found aberrantly at the cortex (arrow) whereas peanut (D’) shows little cortical localization. Anillin and peanut still colocalize (D’’) in ring canals (arrowhead), but these appear somewhat more diffuse than in wild type. (E-F) Male germ cells stained for anillin (green), F- actin (red) and DNA (blue). (E) In early wild-type spermatids, anillin (E) and F-actin (E’) colocalize (E’’) in ring canals. F-actin is also found in the fusome. (F) In contrast, in myt1/Df early spermatids, anillin (F) and F-actin (F’) colocalize (F’’) at the cortex (arrow). Note that the panels in (E) and (F) are from independent experiments and hence are not directly comparable in terms of fluorescence intensity or level of background staining. Bar, 20µm.

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Anillin is phosphorylated

To see if loss of dMyt1 affected anillin protein levels, I performed immunoblots on testis lysates from wild-type and myt1/Df flies (Figure A1.2A). Overall protein levels appeared comparable in wild-type and myt1/Df mutant testes. However, the ratio of slow to fast migrating species in the anillin doublet underwent a dramatic shift in myt1/Df testes. Specifically, the slower migrating species appeared greatly diminished. This suggests that dMyt1 is responsible for post- translational anillin modifications such as phosphorylation.

To test if anillin is phosphorylated in Drosophila, I performed a phosphatase assay on wild-type testis lysates (Figure A1.2B). Homogenization of testes in the presence of various detergents revealed that anillin is largely insoluble in nonionic detergents, a property common to actin-binding proteins (Oliferenko et al., 1999; Seveau et al., 2001). I therefore treated the insoluble pellet recovered after tissue homogenization and centrifugation with lambda phosphatase (see Materials and Methods). Immunoblotting with anti-anillin antibody revealed diminished levels of slower versus faster migrating species in lysates treated with lambda phosphatase, as compared to controls. Thus Drosophila anillin is phosphorylated.

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Figure A1.2. dMyt1 regulates anillin phosphorylation.

(A,B) Immunoblots of wild-type testis lysates probed with anti-anillin antibody. (A) By SDS- PAGE, anillin typically runs as a doublet of ~190 kDa. Note that compared to immunoblots in Figure 3.1, samples shown here were run through a lower percentage SDS-PAGE gel (8% compared to 10%) and for a longer time, making it is easier to see the anillin doublet. Loss of myt1 affects migration of the anillin doublet. Testis lysates from myt1/Df flies contain a lower ratio of slow (asterisk) to fast (double asterisk) migrating species, as compared to wild type. Wild-type and myt1/Df samples contained similar levels of total protein per testis, as determine by Ponceau S staining (not shown). (B) The slower migrating species represents a phosphorylated form of anillin. The ratio of slow to fast migrating anillin species is greatly reduced in lysates treated at 30˚C for 60 minutes with lambda phosphatase, as compared to untreated lysates or to lysates incubated without lambda phosphatase addition.

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Anillin colocalizes with a phosphotyrosine epitope(s) in multiple structures during spermatogenesis

Because lambda phosphatase has activity toward phosphorylated serine, threonine and tyrosine, and since phosphotyrosine (pTyr) epitopes are known to localized to ring canals in both males and females (Robinson et al., 1988; Hime et al., 1996), we considered the possibility that anillin is phosphorylated on tyrosine residue(s). To compare anillin and pTyr localization in these and other male germ cell structures, I performed immunostaining experiments on wild-type testes using anti-anillin antibody and an antibody that recognizes pTyr epitopes. Intriguingly, anillin and pTyr colocalized in a number of structures, including cleavage furrows, ring canals and puncta near elongating nuclei (Figure A1.3).

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Figure A1.3. Anillin colocalizes with phosphotyrosine epitopes in multiple structures during spermatogenesis.

(A-D) Fluorescence (A-C’, D-D’’’) and phase contrast (C’’) micrographs of male germ cells. (A) During late cytokinesis, phosphotyrosine epitopes (pTyr) colocalize with anillin in the cleavage furrow (arrows). (B) Anillin and pTyr remain associated in ring canals (arrowheads) during spermatid elongation. Region bounded by dotted line is magnified two times in the inset. (C, D) Elongated spermatids. (C) pTyr (green) localizes to puncta (arrows) near spermatid nuclei (blue). Note that F-actin (red) containing investment cones have yet to form. (D) pTyr (green) colocalizes with anillin (red) in puncta (arrows) but not in the dense bodies (arrowheads) along the nuclei (blue). Bars (A-C) 20µm and (D) 10µm.

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Discussion

Here, I show for the first time that Drosophila anillin is regulated by dMyt1 and that the hypophosphorylated form of anillin accumulates in myt1 mutant testes. This suggests that anillin is phosphorylated in a cell cycle-dependent manner in flies, and would be consistent with dMyt1 directly phosphorylating anillin. Nonetheless, a direct connection between dMyt1 and anillin appears unlikely, given the known roles for Myt1 in phosphorylating and inhibiting cell cycle- dependent kinases (O'Farrell, 2001; Lindqvist et al., 2009). Loss of Myt1 leads to overactive Cdk1, which, in turn, could lead to misregulation of other kinases, including polo-like kinases, resulting in altered anillin phosphorylation. It should be noted, however, that whereas mammalian Plk1-dependent phosphorylation of Myt1 inhibits its kinase function (Nakajima et al., 2003), Myt1 has not been shown to regulate Plk1. Additional genetic experiments examining single and double myt1 and plo mutant flies will be required to determine whether dMyt1 and Polo act in the same pathway to regulate anillin phosphorylation.

My data suggest that changes in the phosphorylation state of anillin could result in anillin being inappropriately targeted to the cell cortex, where it may bind and recruit or stabilize F- actin. Dephosphorylation of anillin could inhibit interphase nuclear localization by some unknown mechanism. Alternatively, dephosphorylation could promote plasma membrane association by decreasing the negative charge of the protein, thereby potentially stimulating binding to phosphoinositides. Loss of dMyt1 could also result in defects in nuclear structure and/or ring canals in myt1/Df spermatids, which, in turn, could lead to anillin becoming cortical and septins becoming diffuse. Structure-function experiments to determine the mechanism by which anillin localization is regulated and how anillin phosphorylation affects its ability to interact with importins and septins (Silverman-Gavrila et al., 2008) will require identification and mutagenesis of anillin phosphorylation sites.

The finding that Drosophila anillin is phosphorylated agrees with previous observations of S. pombe Mid1p and Xenopus anillin (Sohrmann et al., 1996; Straight et al., 2004). Phosphorylation sites for Drosophila and mammalian anillin have been identified in a number of recent phosphoproteomic screens (Bodenmiller et al., 2007; Chang et al., 2008; Zhai et al., 2008; www.phosphosite.org). Bodenmiller et al. (2007) used Drosophila Kc167 tissue culture cells.

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They identified 28 sites with high confidence, representing 16 unique residues (www.phosphopep.org). Zhai et al. (2008) used embryos, and identified 29 sites with high confidence and 9 unique sites. Five sites overlapped between the two studies. There have been a number of phosphoproteomic studies performed on cultured human cells and data are compiled at the PhosphoSitePlus website. 39 unique sites were identified, including three sites that have been validated in five or more studies. The possibility that anillin is phosphorylated by polo-like kinases is strengthened by the fact that anillin was identified in a screen for PLK1-binding proteins (Lowery et al., 2007). For Drosophila anillin, serine and threonine residues could be tested for phosphorylation by Polo kinase in vitro and the function of this phosphorylation could be examined in rescue experiments.

The data regarding tyrosine phosphorylation of anillin are not yet conclusive. In a tyrosine phosphoproteomic screen performed on S2 cells, one tyrosine-phosphorylated anillin residue was identified in the myosin II interacting region (Chang et al., 2008). Similarly, two tyrosine residues were identified in phosphoproteomic screens of human anillin (www.phosphosite.org). However, these results are from tissue culture cells and may not be relevant to the phosphorylation state of anillin in Drosophila male germ cells. To determine whether anillin is indeed tyrosine-phosphorylated in Drosophila testes, testis extracts prepared in the presence of phosphatase inhibitors could be reciprocally immunoprecipitated and blotted using anti-anillin and anti-pTyr antibodies. In addition, an affinity-tagged version of anillin could be purified from testis extracts in the presence of phosphatase inhibitors and submitted to mass spectrometry for detection of phosphorylated sites (Shou et al., 2002). Finally, rescue constructs carrying mutations (non-phosphorylatable, pseudophosphorylated) in identified tyrosine residues could be used to determine whether the phosphorylation state of this residue is important for anillin function and/or distribution.

In conclusion, the regulation of anillin function over the cell cycle and during development remains poorly understood. Data presented here support the hypothesis that cell cycle kinases regulate anillin phosphorylation. The downstream effects of phosphorylation are still unknown. This is a rich area for future research, as it could lead to identification of residues required to modulate protein-protein and protein-lipid interactions of anillin over the cell cycle.

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