Dissecting the roles of anillin’s interaction with importin β2 and mDia2
by
Anan Chen
A thesis submitted in conformity with requirements for the degree of Master of Science
Department of Biochemistry University of Toronto
© Copyright by Anan Chen, 2015
Abstract
Dissecting the roles of anillin’s interaction with importin β2 and mDia2
Anan Chen
Master of Science
Department of Biochemistry University of Toronto
2015
Abstract
Successful completion of cytokinesis, the final step of cell division, is required to prevent aneuploidy. A vital step in cytokinesis is the formation of an actomyosin ring. Actin filament formation and contraction is key to the ingression of the ring, but the mechanism still remains unclear. Anillin is a scaffolding protein able to cross-link actin with other cytoskeletal elements and regulatory factors of the ring. I sought to examine the roles of the interactions mediated by anillin in cytokinesis. I identified a novel anillin-importin β2 interaction which targets anillin to the nucleus and regulates cell shape in interphase. More importantly, I identified a mechanism by which anillin enhances RhoA dependent activation of formin mDia2. My data suggest a new model whereby anillin acts as an enhancer to help RhoA to fully activate mDia to effectively function in regulating actin organization during ring ingression.
II Acknowledgement
Acknowledgement
I thank my supervisor, Dr. Andrew Wilde, for the guidance, advice and support, as well as my committee members, Dr. Alexander Palazzo and Dr. William Trimble, for their guidance and advice throughout my studies.
I thank my lab members Matt Renshaw and Jinghe Liu for their help and assistance in lab work. I thank Pam Arora and Dr. Christopher McCulloch for their technical assistance and training for the in vitro actin polymerization experiment.
Finally, thank you to The Department of Biochemistry at The University of Toronto. I would like to acknowledge the Canadian Cancer Society for funding this project.
III Table of contents
Contents
Abstract……………………………………………..……………………….…ii Acknowledgement………………………………….……...……………….…iii Table of contents…………………………………………………….………...iv List of figures……………………………………….……………....………..viii List of tables………………………………………………………….……..….x Abbreviation……………………………………………………….…………..xi
1. Introduction……………………………………………………………….…1
1.1 Cytokinesis……………………………………………………………………….……….1 1.1.1 Background introduction………………………………………………………………...1 1.1.2 Cytokinesis and tumorigenesis……………………………………………………….….1 1.1.3 The actomyosin ring……………………………………………………………….…….2 1.1.4 Organization of actomyosin filaments…………………………………………………..5
1.2 Anillin is essential for cytokinesis……………...………………………………………..9 1.2.1 Anillin is a multi-domain scaffolding protein essential for cytokinesis…………………9 1.2.2 C-terminal binding partners of anillin……………………………………………….....13 1.2.3 N-terminal binding partners of anillin……………………………………………….....14 1.2.3.1 F-actin and myosin………………………………………………………………...14 1.2.3.2 mDia2……………………………………………………………………………...14 1.2.3.3 CD2AP…………………………………………………………………………….15 1.2.3.4 The N-terminal NLS……………………………………………………………….15 1.2.3.5 Citron kinase………………………………………………………………………16
1.3 Big questions and strategies……………………………………………………………17
IV Table of contents
Chapter 2. Identification of the anillin nuclear targeting sequence……….18
2.1 Introduction……………………………………………………………………………..18 2.1.1 The nuclear localization of anillin……………………………………………………..18 2.1.2 Objectives and strategies……………………………………………………………….19
2.2 Result…………………………………………………………………………………….19 2.2.1 The NLS lies between N-terminal 1-91aa of anillin…………………………………...19 2.2.2 Sequence analysis reveals potential importin binding site……………..……...……….22 2.2.3 Anillin binds to importin β2 but not importin β1………………………………………25 2.2.4 Mapping the importin β2 binding region on anillin………………………………..…..27 2.2.5 Identification of the importin β2 binding motif of anillin……………………………...27 2.2.6 Importin β2 targets anillin to the nucleus in interphase………………………………..32 2.2.7 The anillin-importin β2 interaction has no significant role in cytokinesis……………..35 2.2.8 The nuclear targeting of anillin contributes to maintain normal cell shape……………39
2.3 Discussion and future direction………………………………………………………..42 2.3.1 The atypical PY-NLS in Nab2 and anillin……………….…………………………….42 2.3.2 Abnormal cell shape caused by cytoplasmic anillin……………..…………………….44
Chapter 3. The role of anillin in cilia function……………………………...45 3.1 Introduction……………………………………………………………………………..45
3.2 Result…………………………………………………………………………………….47 3.2.1 Anillin localizes to the basal body of cilia……………………………………………..47 3.2.2 Anillin RNAi causes defects in cilia length……………………………………………47 3.2.3 The localization of anillin changes during cell differentiation………………………...50 3.2.4 Anillin may not be a truncated isoform in differentiated cells………………………....52 3.2.5 A phospho-mimic mutation does not affect the anillin-importin β2 interaction.……....52
3.3 Discussion and future direction…………………………………………………..……54
V Table of contents
Chapter 4. Anillin enhances RhoA dependent activation of mDia2………56
4.1 Introduction…………………………………………………..………………………...56 4.1.1 Formins and actin nucleation………………………………………………………….56 4.1.2 Regulation of formin activation………………………………………………………..59 4.1.3 Formins in cytokinesis…………………………………………………………………59 4.1.3.1 The pivotal roles of formin in cytokinesis…………………………………………...59 4.1.3.2 The potential role of anillin in mDia2 activation…………………………………….60 4.1.4 Objectives and strategies……………………………………………………………….60
4.2 Result…………………………………………………..………………………………...61 4.2.1 mDia2 has a distinct binding site within region N2 of anilin…………………………..61 4.2.2 Anillin enhances RhoA dependent inhibition of the mDia2 DID-DAD domain interaction…………………………………………………………………………………….65
4.2.3 Anillin does not bind to the NT1-533-CT complex……………………………………...68 4.2.4 Anillin enhances RhoA dependent activation of mDia2 actin polymerization activity………………………………………………………………………………………..71
4.2.4.1 NT1-533 inhibits the actin polymerization activity of CT…………………………..73 4.2.4.2 RhoA partially activates mDia2…………………………………………………...73 4.2.4.3 Anillin has no effect on mDia2 actin polymerization activity…………………….76 4.2.4.4 Anillin enhances RhoA dependent activation of mDia2…………………………..76 4.2.5 The anillin-mDia2 interaction is required for modulating mDia2 activity…………….79
4.3 Discussion and future direction……………………..…………………………………83
5. Summary…………………………………………………………………...88
6. Material and methods……………………………………………………...89
6.1 Materials………………..………………………………………………..……………...89
6.2 cDNA constructs generations…………………………………………..………………90 6.2.1 DNA amplification and isolation………………………………………………………90
VI Table of contents
6.2.2 Cloning in PCR8/GW/TOPO system…………………………………………………..91 6.2.3 Ligation independent cloning (LIC) system……………………………………………91
6.3 Protein expression and purification……………………………………..……………..93 6.3.1 Bacterial transformation………………………………………………………………..93 6.3.2 Protein expression……………………………………………………………………...93 6.3.3 Protein purification…………………………………………………………………….93 6.3.4 Dialysis and storage……………………………………………………………………94 6.3.5 Nucleotide loading……………………………………………………………………..95
6.4 In vitro binding assays…………….……………………………………..……………..96 6.4.1 Anillin and importins binding assays…………………………………………………..96 6.4.2 Anillin and mDia2 binding assays……………………………………………………..97 6.4.3 Anillin, RhoA and mDia2 competition assays…………………………………………97 6.4.4 Importin β2, CD2AP and mDia2 competition assays………………………………….97
6.5 Quantitation of blot intensities……………………………………….………………...99
6.6 In vitro actin polymerization assay …………………………………………………..100
6.6 Immunostaining and microscopy……………………………………………………..103 6.6.1 Cell culture and transfection………………………………………………………….103 6.6.2 Stable cell lines generation and characterization……………………………………..103 6.6.3 siRNA treatment and rescue………………………………………………………….104 6.6.4 Immunofluorescence and microscopy………………………………………………..104
6.7 Statistical analysis………………………………………………………………...... …105
7. Reference……………………………………………………………….....106
Supplementary………………………………………………………………116
VII Table of contents
List of figures
Figure 1.1 Cartoon of cytokinetic events…………………………………...…………………3 Figure 1.2 A direct link between cytokinetic failure, aneuploidy and tumorigenesis…………4 Figure 1.3 Two models of actin filaments organization……………………...………………..7 Figure 1.4 Cartoon of the mechanism of nodes coalescence………………...………………..8 Figure 1.5 Cartoon of anillin domain organization and its interacting partners……………..11 Figure 1.6 The localization of anillin throughout the cell cycle…………...………………...12
Figure 2.1 The NLS lies between 1-91aa on N-terminal of anillin……...…………………...20 Figure 2.2 Quantitation of fluorescence intensities of different constructs………………….21 Figure 2.3 Sequence alignment of anillin………………………………...…………………..23 Figure 2.4 NLSs recognized by importins and the potential NLS of anillin…………………24 Figure 2.5 In vitro binding assays between anillin and importins………...…………………26 Figure 2.6 In vitro binding assays mapping where importin β2 binds to anillin…………….29 Figure 2.7 In vitro binding assays between anillin mutants and importin β2………………..30 Figure 2.8 Quantitation of blot intensities of the pull down group in Fig 2.7………………..31 Figure 2.9 Importin β2 targets anillin to the nucleus in interphase…………………………..33 Figure 2.10 Quantitation of fluorescence intensities of different constructs………………...34 Figure 2.11 Cartoon of anillin RNAi rescue experiments in this study……………………...36 Figure 2.12 Test of expression of anillin in RNAi rescue experiments……………………...37 Figure 2.13 Importin β2 binding to anillin is not required for successful cytokinesis……….38 Figure 2.14 Overexpression of cytoplasmic anillin induces abnormal cell shape…………...40 Figure 2.15 Quantitation of cell shape upon anillin expression……………………………...41 Figure 2.16 Alignment of PY-NLS sequences…………………………………………..…...43
Figure 3.1 Sequence alignment of non-PY motifs of anillin, Nab2 and KIF17……………...46 Figure 3.2 Anillin localizes to cilia in differentiated hTERT RPE-1 cells……...…………...48 Figure 3.3 Anillin RNAi in differentiated hTERT RPE-1 cells……………………...... ……49 Figure 3.4 The localization of anillin is regulated during differentiation……………………51 Figure 3.5 The molecular weight and level of expression of anillin do not change during cell differentiation………………………………………………………………………………...53
VIII Table of contents
Figure 3.6 In vitro binding assays between anillin phospho-mimic mutant and importin β2……………………………………………………………………………………….…….53 Figure 4.1 Three different nucleation machanisms…………………………………………..58 Figure 4.2 Formin domain structure and activation mechanism……………………………..59 Figure 4.3 Sequence alignment of first 100 amino acids of anillin…………………………..63 Figure 4.4 In vitro binding assays mapping the mDia2 binding site on anillin……………...64 Figure 4.5 In vitro binding competition assays between importin β2, CD2AP and mDia2 in binding to anillin……………………………………………………………………………..65 Figure 4.6 Cartoon of proposed models for the roles of anillin in mDia2 activation………..67 Figure 4.7 Cartoon of the in vitro competition assays……………………………………….67 Figure 4.8 In vitro competition assays testing the effect of anillin and RhoA on the interaction between mDia2 fragments NT1-533 and CT……………………………………..……………68
Figure 4.9 In vitro binding assays between anillin and NT1-533 and CT….…………...……..70 Figure 4.10 Anillin binding to the DID domain is dependent on RhoA……………………..71 Figure 4.11 An example of the in vitro pyrene actin polymerization assay…………………73
Figure 4.12 NT1-533 inhibits the actin polymerization activity of CT…………………....…...75 Figure 4.13 RhoA partially activates mDia2…………………………………………………76 Figure 4.14 Anillin does not activate mDia2 actin polymerization activity…………………78 Figure 4.15 Anillin enhances the activation effect of RhoA on mDia2……………………...79 Figure 4.16 In vitro binding assays reveal anillin residues that contribute to mDia2 binding………………………………………………………………………………………..81 Figure 4.17 Quantitation of blot intensities of the pull down groups in Fig 4.16……………82 Figure 4.18 The anillin-mDia2 interaction is required for modulating mDia2 activity……...83 Figure 4.19 A new model of cellular formin activation and a list of DID-binding ligands………………………………………………………………………………………..87
Figure 6.1 Graphical representation of the methods to analyze the in vitro pyrene actin polymerization data…………………………………………………………………………102
IX Lists of figures and tables
List of Tables (Supplementary)
Table 6.1 Kits used in this work…………………………………………………………………….116 Table 6.2 Oligonucleotides used to generate wild-type constructs...……………………………….117 Table 6.3 Oligonucleotides used to generate mutant constructs………………………..…………..119
X Abbreviation
Abbreviation
AH (domain) Anillin homology (domain) CLS Cilia localization sequence C-terminal Carboxyl terminal CT C-terminal region of mDia2 containing amino acid residues 560-1193 DAD Diaphanous autoregulatory domain DID Diaphanous inhibitory domain DRF Diaphanous-related formin F-actin Filamentous actin FH (domain) Formin homology (domain) GBD G-protein binding domain GFP Green fluorescence protein GST Glutathione S-transferase ICB Intercellular bridge MBP Maltose binding protein
NT1-533 N-terminal region of mDia2 containing amino acid residues 1-533
NT89-533 N-terminal region of mDia2 containing amino acid residues 89-533 NTR Nuclear transport receptor N-terminal Amino terminal NLS Nuclear localization sequence PCR Polymerase chain reaction RBD Rho binding domain SH3 (domain) SRC homology 3 (domain) TIRFM Total internal reflection fluorescence microscopy UTR Untranslated region
XI Introduction 1.1 Cytokinesis
1.Introduction
1.1 Cytokinesis
1.1.1 Background introduction
Cytokinesis is the final step in mitosis when a cell physically divides into two new daughter cells (Fededa and Gerlich 2012). In animal cells, cytokinesis initially begins with the reorganization of the mitotic spindle and the specification of the division plane between the segregated chromosomes (Fig 1.1A). The spatial signal that determines the location of the division plane is transmitted through the RhoA pathway, which stimulates actomyosin filament formation and leads to the assembly of an equatorial contractile ring (Fig 1.1B). As the ring is tethered to the plasma membrane, when it constricts, the membrane ingresses (Eggert et al. 2006). As the constriction progresses, the central region of the anaphase spindle is remodeled to form a narrow intercellular bridge between the nascent daughter cells (Fig 1.1C). Subsequently, the final abscission proceeds by removal of cytoskeletal structures from the intercellular bridge followed by the fission of the plasma membrane, that leads to the final separation of the daughter cells (Renshaw et al. 2014) (Fig 1.1D). One of the important consequences of cytokinesis is to generate two new daughter cells each with one intact copy of the genome to secure the faithful inheritance of genetic materials. Disruption of cytokinesis leads to mitosis failure, aneuploidy and tumorigenesis (Ganem et al. 2007).
1.1.2 Cytokinesis and tumorigenesis
It is vital that cytokinesis happens at the right time and place within the cell to generate diploid daughter cells each with a complete copy of the genome. If cytokinesis is disrupted in any of the stages depicted in Fig 1.1, the consequence is that although the chromosomes are still segregated, mitosis fails and results in a tetraploid cell (Fig 1.2A). Instead of having a single centrosome the tetraploid cell has two centrosomes and multiple copies of the genome. Subsequently, the next round of division will randomize the splitting of genome copies into daughter cells. As a result, the daughter cells are aneuploid, a hallmark of cancer. Indeed,
1 Introduction 1.1 Cytokinesis people have demonstrated a direct link between tetraploid cells and tumor formation. Injection of tetraploid cells (resulting from cytokinesis failure) into mice leads to tumorigenesis. In contrast, injection of the diploid cells does not (Fig 1.2B). These results suggest a direct link between cytokinetic failure, aneuploidy and tumorigenesis (Fujiwara et al. 2005).
1.1.3 The actomyosin ring
A core structure that drives furrow ingression is the contractile ring, which is also termed the actomyosin ring (Schroeder 1973). The primary component of the ring is the actomyosin cytoskeleton (Mabuchi et al. 1988). It also consists of additional cytokinetic factors like anillin, formins, and septins. The ring is highly dynamic with fast turn over rates for actin and myosin (Yumura 2001; Murthy and Wadsworth 2005). During early anaphase the ring is assembled at the equatorial cortex and tethered to the membrane. It is still not clear how the ring is tethered to the membrane. Studies have suggested multiple factors are involved in this process, including anillin which can interact with actomyosin (Field and Alberts 1995; Straight et al. 2005), the septin cytoskeleton (Kinoshita et al. 2002; Liu et al. 2012) and the membrane via the lipid PIP2 (Liu et al. 2012). Therefore anillin may contribute to anchor the ring at the membrane. The constriction of the ring progressively pulls the membrane inward to drive the furrow ingression (Fig 1.1).
2 Introduction 1.1 Cytokinesis
Fig 1.1
Cartoon of cytokinetic events (A) In early anaphase, the mitotic spindle reorganizes to form a central spindle which specifies the division plane between the segregating chromosomes. (B) During anaphase, active RhoA enriches at the membrane furrow and directs contractile ring assembly. The ring is tethered to the membrane and constricts to ingress the membrane inward. (C) In late anaphase and telophase, the membrane furrow continues to ingress and the central region of the anaphase spindle is remodeled to form a narrow intercellular bridge (ICB) between the daughter cells. (D) Final abscission leads to the separation of two daughter cells.
3 Introduction 1.1 Cytokinesis
Fig 1.2
A direct link between cytokinetic failure, aneuploidy and tumorigenesis. Cells experiencing normal cytokinesis generate two diploid daughter cells each with one complete copy of genome. Disruption in actomyosin ring assembly, furrow ingression or intercellular bridge formation can lead to cytokinetic failure which results in tetraploid cells. Injection of tetraploid cells into mice leads to tumor formation, whereas when injection of diploid cells does not induce tumor formation (Fujiwara et al. 2005).
4 Introduction 1.1 Cytokinesis
1.1.4. Organization of actomyosin filaments
Previously, studies showed that the translocation of actin filaments by bipolar myosin filaments is the force generating mechanism for ring constriction (Noguchi et al 2001; Matsumura 2005). The key question of how the actomyosin filaments are organized and exert force is unclear. An early dominant model to describe the actomyosin filaments organization was the purse-string model (Schroeder 1968, Schroeder 1972), which built upon the idea of how actomyosin filaments worked in muscle where parallel filaments slide across each other and generate contraction (Fig 1.3A). Yet there is little structural evidence for the alignment of cortical actin fibers in this orientation (Maupin and Pollard 1986; Mabuchi et al. 1988). In electron microscopy studies the majority of cortical actin fibers in both yeast and mammalian cells are randomly oriented (Kamasaki et al. 2007) (Fig 1.3B). These data suggest the purse-string data does not reasonably explain cortical actomyosin ring organization.
An alternative ‘node coalescence’ model from studies in fission yeast may better describe the organization of the ring (Fig 1.3C) (Wu et al. 2003). During the initiation of cytokinesis, the anillin-like protein Mid1p was first released from the nucleus and established a broad band of small dots or nodes in the cortex near the nucleus (Fig 1.3C, D) (Wu et al. 2003). Mid1p is the initial constituent of the macromolecular node structure and can recruit other factors including myosin II, formins and formin activators that direct actin filament assembly (Vavylonis et al. 2008, Laporte D et al. 2011). Actin filaments grow from nodes and are captured by myosin filaments from adjacent nodes. Myosin is proposed to pull the nodes together thereby driving coalescence (Fig 1.4). Subsequently, these nodes coalesced laterally into a compact ring that aggregated further to drive the furrow ingression (Fig 1.3C, D) (Wu et al. 2003). In the node model, the actin filaments are initially generated from different nodes and then assembled together during nodes coalescence, which fits with the random actin filaments organization observed in the ring. Additional support for this model comes from mathematical modeling in C.elegans embryos analyzing the rates of furrow ingression. This study indicated that there were a series of defined constriction units that coalesce. When coupled with actin de-polymerization, they drive membrane ingression (Carvalho et al. 2009). The key question is, does this node model apply to metazoan cells? Furthermore, how are actin filaments generated and organizated to regulate cytokinesis in metazoan cells?
5 Introduction 1.1 Cytokinesis
Despite these different models for actomyosin filament organization, it is still not clear how myosin II contributes to generate the contraction force in the network of actomyosin filament. A recent study showed that a non-muscle myosin II mutant with impaired actin-activated ATPase activity and an inability to translocate actin filaments had no effect on cytokinesis (Ma et al. 2012). This data suggest that the role of non-muscle myosin II in cytokinesis may not depend on its enzymatic motor activity in translocation of actin filaments, but rather that non-muscle myosin II crosslinks actin filaments to maintain tension during actomyosin ring constriction (Ma et al. 2012).
6 Introduction 1.1 Cytokinesis
Fig 1.3
Two models of actin filaments organization. (A) In the purse-string model, parallel filaments slide across each other to generate force. (B) EM image showing the random orientation of cortex actin filaments in yeast. (C) In the ‘node’ coalescence model, actin filaments are initially organized from different nodes and assembled into a ring structure when nodes coalesce. (D) An image showing node distribution at the central region of plasma membrane (top red arrow) and coalescence into an actomyosin ring structure (bottom red arrow).
7 Introduction 1.1 Cytokinesis
Fig 1.4
Cartoon of the mechanism of node coalescence. Mid1p (Anillin homolog in yeast) establishes nodes by recruiting myosin II motor, formins and formin activators. Actin filaments are organized from individual nodes and captured by myosin II motors from adjacent nodes. The interaction between myosin and actin filaments is proposed to pull these nodes together thereby driving coalescence.
8 Introduction 1.2 Anillin is an essential cytokinetic organizer
1.2 Anillin is an essential cytokinetic organizer
1.2.1 Anillin is a multi-domain scaffolding protein essential for cytokinesis
Anillin is a multi-domain scaffolding protein essential for the successful completion of cytokinesis (Piekny and Maddox 2010). Anillin is reported to bind many cytokinetic factors (Fig 1.5), including the actomyosin cytoskeleton (Miller et al. 1989; Straight et al 2005; Piekny and Maddox 2010) and the septin cytoskeleton (Kinoshita 2002; D’ Avino et al. 2009; Liu et al. 2012). In addition, the Wilde lab has shown that anillin can bind to phosphatidylinositol 4,5-biphosphate (PIP2) which concentrates at the furrow (Liu et al 2012). Therefore, anillin has the capacity to contribute to tethering the actomyosin ring to the plasma membrane. With its multi-domain interacting capacity, anillin is an excellent candidate for organizing different cytokinetic factors and facilitating the transmission of different cellular signals back and forth between the plasma membrane and the cytoskeleton throughout cytokinesis.
In S. pombe, anillin-like protein Mid1p is the initial builder of the nodes that coalesce to form contractile ring (Wu et al. 2006). Loss of Mid1p function disrupts either ring formation or ring positioning (Huang et al. 2008). In metazoan cells, during early anaphase anillin is proposed to crosslink F-actin, myosin and septin cytoskeleton at the equatorial cortex (Miller et al. 1989; Straight et al. 2005; Liu et al. 2012). During constriction of the ring, anillin also helps in balancing the contractility of the polar cortex to prevent shape oscillations and unbalanced division (Sedzinski et al. 2011). In the late-phases of cytokinesis anillin participates in the formation of the intercellular bridge (Renshaw et al. 2014). In Drosophila S2 cells, anillin localizes to the furrow in early anaphase of cytokinesis. However depletion of anillin reveals roles only in furrow stabilization and ICB formation during late anaphase and telophase (Echard et al. 2004; Hickson and O’Farrell 2008). Studies have shown a unique Rho-dependent input allows stable filamentous structures containing anillin and septins to form at the equatorial cortex during the time of furrowing, which is independent of F-actin and myosin (Hickson and O’Farrell 2008).
9 Introduction 1.2 Anillin is an essential cytokinetic organizer
Anillin’s subcellular localization changes throughout the cell cycle (Field and Alberts 1995; Oegema et al. 2000; Liu et al. 2012) (Fig 1.6). Anillin is targeted to nucleoplasm during interphase in Drosophila and human cultured cells, and in fission yeast (Field and Alberts 1995; Oegema et al. 2000; Wu et al. 2003). Upon mitotic entry, the nuclear envelope breaks down and anillin is released into cytoplasm where it re-localizes to the membrane furrow (Oegema et al. 2000; Liu et al. 2012). Anillin remains at the furrow when the intercellular bridge matures and is gradually depleted from the ICB after relocating to the sites of microtubule constriction (Renshaw et al. 2014). The progressive loss of anillin is septin-dependent (Renshaw et al. 2014). During mitotic exit, anillin is imported back to the nucleus and its level drops (Field and Alberts 1995; Zhao and Fang 2005). Anillin is a substrate of the anaphase-promoting complex/cyclosome (APC/C) and contains a destruction box (D-box) at amino acid residues 41-44 (Zhao and Fang 2005). Mutations in this D-box disrupt the recognition by APC/C and the level of the mutant does not decrease during the M/G1 transition (Zhao and Fang 2005).
10 Introduction 1.2 Anillin is an essential cytokinetic organizer
Fig 1.5
Cartoon of anillin domain organization and its putative interacting partners. Anillin is a multi-domain protein that binds to different factors and plays pivotal roles during cytokinesis. All llisted binding factors will be discussed in chapter 1.2.1, 1.2.2 and 1.2.3. * marked interactions are the focus of my study.
11 Introduction 1.2 Anillin is an essential cytokinetic organizer
Fig 1.6
The localization of anillin throughout the cell cycle. Anillin is targeted to the nucleus in interphase. In mitosis when the nuclear envelope breaks down, anillin is released from the nucleus to the cytoplasm. In early anaphase anillin is concentrated at the membrane furrow and participates in furrow ingression. In late anaphase and telophase, anillin localizes to the intercellular bridge untill the final separation of the daughter cells. (Anillin in green; microtubules in red; DNA in blue)
12 Introduction 1.2 Anillin is an essential cytokinetic organizer
1.2.2 C-terminal of anillin and its binding partner
The C-terminal region of anillin contains the anillin homology (AH) domain and the pleckstrin homology (PH) domain where multiple factors bind (Fig 1.5). In mammalian cells, anillin interacts with septins through the PH domain plus part of the upstream sequence (Kinoshita et al 2002; Silverman-Gavrilla et al. 2008; Liu et al. 2012). The association of anillin and septins contributes to the localization of both to the cortex and the cytokinetic ring in Drosophila and human cells (Field et al. 2005; Liu et al 2012). In addition, the PH domain of anillin also bound to phosphatidylinositol 4,5-biophosphate (PIP2), a lipid that concentrates in the membrane of the furrow. The anillin-PIP2 interaction targets anillin to the furrow (Fig 1.6) (Liu et al. 2012). Therefore suggests anillin has the capacity to contribute to anchoring the actomyosin ring at the membrane furrow.
Anillin directly interacts with RhoA through the AH domain (Fig 1.5) (Liu et al. 2012; Piekny et al. 2008). In Drosophila S2 cells, this interaction was required for the equatorial cortex localization of anillin (Hickson and O’Farrel 2008). In human cells, their interaction was implicated in the generation and stabilization of active RhoA in the division plane (Piekny and Glotzer 2008). Anillin also interacts with the RhoA activator Ect2 (Fig 1.5) (Frenette et al. 2012). The anillin-Ect2 interaction was proposed to stabilize active RhoA localization at the division plane (Piekny and Maddox 2010) and was also implicated in the cortical localization of central spindle proteins (Frenette et al. 2012).
In Drosophila, anillin may participate in division plane positioning via the binding between its C-terminus and the central spindle protein RacGAP50C (Fig 1.5) (D’Avino et al. 2008, Gregory et al. 2008). In the absence of anillin, the spindle-associated RacGAP lost its association with the equatorial cortex and cytokinesis failed (Gregory et al. 2008). In addition, anillin is suggested to interact with microtubules, although this interaction appears to be weak (Fig 1.5) (Field and Alberts 1995; Van et al. 2014). The anillin- microtubule interaction may also contribute to the specification of the division plane (Van et al. 2014).
The interactions between the C-terminal region of Drosophila anillin and importins (α and β1) was shown to regulate septin recruitment to the pseudocleavage furrow in Drosophila
13 Introduction 1.2 Anillin is an essential cytokinetic organizer embryos via the RanGTP pathway (Fig 1.6) (Silverman-Gavrila et al. 2008). Interestingly, this C-terminal importin α/β1 binding site was not responsible for the nuclear targeting of anillin in interphase (Silverman-Gavrila et al. 2008). Instead, a second undefined NLS in N-terminal region of anillin was the predominant site for nuclear targeting (Oegema et al. 2000).
1.2.3 N-terminal of anillin and its binding partners
The N-terminal region of anillin has multiple binding partners including actin (Miller et al 1989), myosin (Straight et al. 2005), mDia2 (Watanabe et al. 2008), CD2AP (Monzo et al. 2005) and citron kinase (Gai et al. 2011). The roles of N-terminal region of anillin have been implicated in different stages of cytokinesis (Gai et al. 2011; Watanabe et al. 2008; Straight et al. 2005; Oegema et al. 2000), but the role of specific interaction has not been determined. The binding partners are described in 1.2.3.1-1.2.3.5.
1.2.3.1 F-actin and myosin
Anillin binds to actin and myosin (Field and Alberts 1995; Straight et al. 2005). The actin binding site lies in the conserved residues 258-340aa in Drosophila (Field and Alberts 1995) and residues 231-454aa in Human (Oegema et al. 2000). Myosin II binds to a conserved domain containing residues 142-254aa in Xenopus (Straight et al. 2005) (Fig 1.5). Anillin specifically co-localizes with contractile ring-related F-actin and plays roles in actomyosin ring organization at the equatorial cortex (Field and Alberts 1995; Kinoshita et al. 2002). In addition, the interaction between anillin and actomyosin filament has been implicated in modulating contractility (Werner and Glotzer 2008). In Drosophila, anillin depletion causes later defects in furrow destabilization and abnormal ICB formation that results in cytokinesis failure (Echard et al. 2004; Hickson and O’Farrell 2008; Goldbach et al. 2010). As described in chapter 1.1.3 and Fig 1.5, in S.pombe, Mid1p (the anillin homolog) recruits myosin to the pre-ring nodes that capture F-actin filaments and organize the contractile ring (Wu et al. 2006). However, the role of anillin interaction with actin and myosin has never been directly tested.
14 Introduction 1.2 Anillin is an essential cytokinetic organizer
1.2.3.2 mDia2
Formins are actin nucleators (Pruyne et al. 2002; Sagot et al. 2002). The Diaphanous formin mDia2 belongs to the diaphanous-related formins (DRFs) family that nucleates actin filaments in metazoan cells (Goode and Eck 2007). The very N-terminal 1-91aa of anillin directly interact with mDia2 (Watanabe et al. 2008) (Fig 1.5). This interaction contributes to the positioning of mDia2 at the furrow (Watanabe et al. 2008). The presence of anillin is essential for mDia2 furrow localization (Watanabe et al. 2010). As the upstream effector of mDia2, RhoA is also proposed to recruit mDia2 to the furrow and activate it (Watanabe et al. 2008). However, there is mounting evidence that RhoA does not fully activate formins, suggesting other factors are involved in modulating mDia2 activity (Li and Higgs 2005; Maiti et al. 2012).
1.2.3.3 CD2AP
CD2AP (cluster of differentiation 2-associated protein) plays roles in membrane trafficking and actin remodeling (Lehtonen et al. 2002; Cormont et al. 2003). Human anillin directly interacts with CD2AP via a conserved SH3 domain binding site from amino acid residues P27 to R32 (Monzo et al. 2005) (Fig 1.5). Drosophila anillin also interacts with Cindr (CD2AP orthologue) (Haglund et al 2010). CD2AP has been implicated in the stabilization of the intercellular bridge and in the final abscission event (Monzo et al. 2005). However the role of the anillin-CD2AP interaction in cytokinesis has not been directly tested (Monzo et al. 2005).
1.2.3.4 The N-terminal NLS
Anillin is targeted to the nucleus during interphase (Field and Alberts 1995; Oegema et al. 2000). Importins are nuclear transport receptors (NTRs) that mediate cargo nuclear localization via binding to the nuclear localization sequences (NLSs) of the cargo (Chook and Blobel 2001). In Drosophila embryos, anillin was reported to bind to importin β1 via its C-terminus (Silverman-Gavrilla et al. 2008). This interaction regulates septin recruitment to the pseudocleavage furrow through the RanGTP pathway, yet it does not contribute to the nuclear targeting of anillin (Silverman-Gavrilla et al. 2008). A NLS that targets anillin to the
15 Introduction 1.2 Anillin is an essential cytokinetic organizer nucleus was mapped to the first 1-230aa of human anillin (Oegema et al. 2000), a region that overlaps with the mDia2, CD2AP and myosin binding sites (Fig 1.5). It raises the question of a role for the RanGTP pathway in regulating the mitotic function of anillin and anillin binding partners (mDia2, CD2AP) via affecting the undefined NTR-NLS interaction. Another interesting question is if the nuclear targeting of anillin contributes to the cytokinetic roles of anillin, or other general cellular process. One possibility is to prevent any disrupting effects caused by excessive cytoplasmic anillin in interphase. After all, as a cytoskeleton binding protein (Straight et al. 2005; Kinoshita 2002; Field and Alberts 1995) anillin should be compartmentalized in interphase from the global cytoskeletal structure before it is properly targeted and becomes active.
1.2.3.5 Citron kinase
Citron kinase is required for cytokinesis and is considered a cytokinesis-specific effector of active RhoA (Naim et al. 2004). In vitro, citron kinase phosphorylates regulatory myosin light chain (MLC) of myosin II (Yamashiro et al. 2003). However it is still unclear if citron kinase is involved in regulating MLC phosphorylation in vivo (Yamashiro et al. 2003; Matsumura et al. 2005; Dean and Spudich 2006). Citron kinase has been implicated in retaining both anillin and RhoA at the intercellular bridge. Anillin interacts with citron kinase via the N-terminal first 1-250aa (Gai et al. 2011) (Fig 1.5). In Drosophila, citron kinase deletion results in abnormal ICB formation and delayed abscission (Naim et al. 2004). It has been implicated in promoting ICB stability through anillin and RhoA (Gai et al. 2011). .
16 Introduction 1.3 Big questions and strategies
1.3 Big questions and strategies
Anillin binds to different cytokinetic factors and its localization changes throughout cytokinesis, suggesting that it plays multiple roles (Renshaw et al. 2014; Liu et al. 2012; Piekny and Glotzer 2008). I’m focusing on the N-terminal region of anillin, which binds to multiple factors and has been implicated in different stages of cytokinesis (Gai et al. 2011; Watanabe et al. 2008; Monzo et al. 2005; Straight et al. 2005; Oegema et al. 2000). Previous studies have deleted regions at the N-terminus of anillin that include multiple binding sites and the roles of individual interactions have not been directly tested. An important question is what is the biological role of each individual interaction? Moreover, what are the interrelationships of these interactions and are there any regulatory mechanisms of these interactions on each other?
By establishing an anillin mutant that specifically blocked septin binding, the Wilde lab was able to characterize the pivotal role of the anillin-septin interaction in recruiting septins to the furrow (Liu et al. 2012). I used this strategy to generate mutations in anillin that specifically disrupt individual interactions and analyzed the role of individual interactions in cytokinesis.
In the following chapters, I first identified a novel interaction between anillin and importin β2 which targets anillin to the nucleus and contributes to maintaining normal cell shape in interphase. Secondly, I discovered the ciliary targeting of anillin and analyzed the role of anillin in ciliogenesis. Thirdly, I identified a novel formin activation mechanism where anillin enhances RhoA activation on mDia2 actin polymerization activity.
17 Chapter 2 Identification of the anillin nuclear targeting sequence
Chapter 2. Identification of the anillin nuclear targeting sequence
2.1 Introduction
2.1.1 The nuclear localization of anillin
Many proteins with mitotic functions are targeted to the nucleus during interphase either to perform additional interphase functions (such as MgcRacGAP, Lagana et al. 2010) or to prevent them from disrupting the interphase cytosol (such as TPX2, Trieselmann et al. 2003). Therefore understanding how and why proteins with a mitotic function are targeted to the nucleus is important in understanding the breadth of their function within the cell. In addition, some mitotic proteins that have nuclear localization signals continue to be regulated by the mitotic Ran pathway that directs the spatial organization of the mitotic cell (Wiese et al. 2001; Nachury et al. 2001; Trieselmann et al. 2003; Tsai et al. 2003; Zonis and Wilde 2011).
Anillin localizes to the plasma membrane, the actomyosin ring and the intercellular bridge during cytokinesis (Piekny and Glotzer 2008; Renshaw et al. 2014). However in interphase anillin is targeted to the nucleus (Field and Alberts 1995; Oegema et al. 2000). A previous study found that the C-terminus of anillin bound to nuclear transport receptors (NTRs) (importin α and β1) and binding regulated anillin-septin interaction at the pseudocleavage furrow in Drosophila embryos (Silverman-Gavrilla et al. 2008). However, this interaction was not responsible for targeting anillin to the nucleus. There was evidence suggesting the NLS resided in first 1-230 amino acids of anillin (Oegema et al. 2000). The mechanism by which anillin is targeted to the nucleus was unknown when I began this project, nor was the relationship of nuclear targeting to cytokinesis understood.
18 Chapter 2 Identification of the anillin nuclear targeting sequence
2.1.2 Objectives and strategies
Identifying the anillin-NTR interaction will reveal how the localization of anillin is regulated throughout the cell cycle. More importantly, it will help us understand whether the nuclear targeting contributes to the cytokinetic roles of anillin. To determine the N-terminal NLS that targets anillin to the nucleus, I will map the NLS in the N-terminus of anillin and identify the NTR that interacts with anillin and targets it to the nucleus. By generating point mutants that block NTR binding and nuclear targeting, my research will reveal more broadly the cellular roles of the anillin-NTR interaction.
2.2 Results
2.2.1 The NLS lies between N-terminal 1-91aa of anillin
To map which domain of anillin contained the NLS, I first fused different fragments of anillin including 1-300, 91-450, 600-1124 and 1-1124 (full length, FL) to GFP (Fig 2.1A). The constructs were transiently expressed in HeLa cells, fixed and then their subcellular localizations determined by microscopy. An anillin fragment containing the C-terminal half of the protein, amino acids 600-1124, did not localize to the nucleus (Fig 2.1B). In contrast, an anillin fragment containing amino acids 1-300 localized to the nucleus in interphase cells (Fig 2.1B). These data suggest the NLS lies in the amino terminus of anillin, specifically in the first 300 amino acids, which is consistent with previous observations (Oegema et al. 2000). Moreover, a fragment of anillin lacking the first N-terminal 91 amino acids localized to the cytoplasm rather than the nucleus (Fig 2.1B), demonstrating that NLS lies within the first 91 amino acids on N-terminal of anillin.
The difference in construct localization was further analyzed by comparing the ratio of fluorescence intensity of GFP signals (nucleus to the cytoplasm) (Fig 2.2). Constructs that localized to the cytoplasm rather than the nucleus (91-450 and 600-1124) had significantly lower ratio of fluorescence intensity than constructs that localized to the nucleus (1-300 and 1-1124) (Fig 2.2).
19 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.1
The NLS lies between 1-91aa on N-terminal of anillin. (A) Cartoon of anillin fragments fused to GFP and expressed in cells. (B) Immunofluorescence images showing the localization of different anillin fragments in interphase cells. Construct localization was visualized by GFP signal. Scale bars=10µm.
20 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.2
Quantitation of fluorescence intensities of different constructs. Constructs that did not localize to the nucleus had significantly lower ratio of fluorescence intensities (nucleus to cytoplasm) than the full length wild-type construct. More than 100 cells were quantitated in each group (n=3). Error bars indicate ± SEM.
21 Chapter 2 Identification of the anillin nuclear targeting sequence
2.2.2 Sequence analysis reveals potential importin binding site
To determine where the NLS may reside in the first 91 amino acids of anillin I first compared anillin sequences from different vertebrate species (Fig 2.3). This region is evolutionarily conserved and can be further broken into 4 distinct conserved regions that I termed N1, N2, N3, and N4 (Fig 2.3). This region binds to multiple factors and is critical for the cytokinetic functions of anillin. It has been reported to bind to mDia2 (Watanabe et al. 2008). It also has the conserved residues 27-32 within N1 that bind to the first 3 SH3 domains of CD2AP (Monzo et al. 2005). However, the rest of the interaction sites on N2, N3 and N4 are unknown.
The most common mechanism for of NLSs-mediated cargo nuclear localization is through binding to nuclear transport receptors (NTRs) such as importins (Chook and Blobel 2001). Importin-binding NLSs can vary, but most of them contain a core of basic amino acids residues (Chook et al. 2011). The classical importin α/β1-binding NLSs are either bipartite or monopartite motifs (Fig 2.4) (Pemberton et al. 1998; Cingolani et al. 1999; Lee et al. 2003). After a careful analysis of the N1-N4 region, I found a conserved basic patch of residues, KKR (68-70), within N3 that could be the binding site of importin β1 (Fig 2.4).
22 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.3
Sequence alignment of anillin. Cross-species sequence alignment of the first 1-100 amino acids of anillin showing highly evolutionary conserved regions divided as N1-N4. The darker the color, the greater the sequence conservation. The binding region of mDia2 lies within the amino acid residues 1-91. The potential importins binding motifs are marked with asterisks (also indicated in Fig 2.4). The CD2AP binding motifs are marked in triangles.
23 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.4
NLSs recognized by importins and the potential NLS of anillin. (A) Classical bipartite or monopartite NLSs that bind to importin α/β1. Non-classical NLSs that have basic cluster at the upstream and the (-PY-) motif at the downstream bound to importin β2 (Φ=hydrophobic residue). (B) The potential NLS of anillin (human) contains a basic patch of residues at amino acid 68-70 (marked with asterisk at left). The (PV) motif downstream (marked with asterisk at right) the basic patch bares close resemblance to a basic-PY motif that binds to importin β2.
24 Chapter 2 Identification of the anillin nuclear targeting sequence
2.2.3 Anillin binds to importin β2 but not importin β1
To determine the mechanism of anillin targeting to the nucleus I analyzed the direct binding of recombinant anillin fragments to NTRs. The classical bipartite or monopartite NLSs bind to either importin β1 or a complex with importin β1 and importin α (Fig 2.4) (Chook et al. 2011). A fragment of anillin containing the first N-terminal 151 amino acids was generated and fused to maltose-binding protein (MBP) to test the binding with importin β1 in vitro. Proteins were mixed, re-isolated on amylose resin and analyzed by SDS-PAGE and Western-blotting using an anti-6xHis antibody. Surprisingly, the anillin 1-151 fragment did not bind to importin β1 or a complex of importin β1 and α (Fig 2.5A), suggesting another NTR must be involved in targeting anillin to the nucleus.
I re-analyzed the anillin sequence and discovered a conserved region in N4 contained a conserved proline (amino acid 88) that was preceeded by a basic residue (Fig 2.3, Fig 2.4). Although the proline is not followed by a tyrosine, the combination of conserved regions N3 and N4 bears close resemblance to a basic patch-PY motif that binds importin β2 (Lee et al. 2006) (Fig 2.4). The substrate sequence of importin β2 consists of an upstream basic patch of residues with the downstream (PY) motif (also termed as PY-NLS) (Fig 2.4) (Lee et al. 2006). I hypothesized the PV (87-88) at N4 combined with the KKR (68-70) at N3 could be an importin β2 binding motif (Fig 2.4). To test this model, I generated recombinant 6xHis tagged importin β2 and tested its ability to bind to the anillin 1-151 fragment. Interestingly, 6x His tagged importin β2 bound to anillin (Fig 2.5A).
Importins bind cargo in the cytoplasm where there are low levels of RanGTP. In the nucleus, where the level of RanGTP is high, RanGTP binds to importins and releases their cargo (Gorlich and Kutay 1999). In the presence of RanGTP, importin β2 does not bind to anillin (Fig 2.5B), suggesting the anillin-importin β2 interaction reflects a physiological NTR-cargo interaction. These data suggest anillin is targeted to the nucleus in an importin β2 dependent manner.
25 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.5
In vitro binding assays between anillin and importins. (A) Western-blotting analysis showing co-purifying 6xHis-importins by MBP/MBP-anillin pull-down assays. MBP/MBP-anillin was pulled down and co-purifying 6xHis-importins were analyzed. (B) Western-blotting analysis testing co-purifying importin β2 by MBP-anillin pull-down assays in the presence or absence of RanGTP. The band of anillin is indicated by the arrow. (Anln=anillin)
26 Chapter 2 Identification of the anillin nuclear targeting sequence
2.2.4 Mapping the importin β2 binding region on anillin
To define the importin β2 binding site on anillin I generated anillin fragments 1-57, 1-91, 1-151, 1-300, 57-151, 22-151 and fused them to MBP to analyze their binding to importin β2 (Fig 2.6A). Results showed anillin fragments containing amino acid residues 57-91 bound to importin β2 (Fig 2.6B). These data suggest the sequence stretching from KKR (68-70) to the downstream PV (87-88) in regions N3 and N4 that have similarity to importin β2 binding motifs are indeed the sites of importin β2 binding to anillin.
2.2.5 Identification of the importin β2 binding motif on anillin
Based on the results from the sequence analysis in chapter 2.1.2 and mapping assays in 2.1.4, I hypothesized that KKR (68-70) combined with the downstream PV (87-88) could be a PY-NLS like motif that binds to importin β2. PY-NLSs were typically defined as a patch of hydrophobic or basic residues upstream of the (R/H/K)-x(2-5)-(PY) motif (Fig 2.4) (Lee et al. 2006; Xu et al. 2010). Yet previous studies have revealed only two PY-NLSs in which the core proline is flanked by other hydrophobic residues instead of tyrosine (Suel et al. 2008; Soniat et al. 2013). In one case where (PY) is replaced by (PL) in the PY-NLS of yeast Nab2, crystallography analysis identified several residues neighboring the central proline (including the leucine) that contributed to importin β2 binding (Soniat et al. 2013). To further understand how importin β2 binds to the cargo it is necessary to understand which residues in the (PV) motif of anillin contribute to importin β2 binding. Further examination of the cross species sequence alignments of anillin identified 3 residues directly upstream of the PV (87-88) that were conserved across vertebrates, E83, N84 and Q86 (Fig 2.3), which may contribute to importin β2 binding.
To determine if the proposed motifs and their neighboring residues in N3 and N4 of anillin actually contribute to importin β2 binding, I generated anillin fragments 1-151 with a series of point mutations including KKR (68-70)-AAA, E83A, N84A, Q86A, PV (87-88)-AA and a mutant containing both KKR (68-70)-AAA and PV (87-88)-AA. These anillin fragments were fused to MBP and analyzed for their ability to bind to importin β2 in vitro and analyzed by Western-blotting (Fig 2.7A-C), which were further quantified (Fig 2.8). Mutation of PV
27 Chapter 2 Identification of the anillin nuclear targeting sequence
(87-88) to alanines reduced the binding to importin β2 by 52±7.5% (p<0.05) (Fig 2.7B; Fig 2.8). In contrast, mutation of the basic patch residues KKR (68-70) to alanines almost completely abrogated the binding to importin β2, reducing binding by 93±1.5% (p<0.05) (Fig 2.7B; Fig 2.8). Likewise, the mutant with the combined mutations of KKR (68-70)-AAA and PV (87-88)-AA severely reduced the binding with importin β2 by 97±1.0% (p<0.05) (Fig 2.7B; Fig2.8). Mutation of E83, N84 and Q86 each to alanine revealed that only mutation of Q86 significantly reduced the binding with importin β2 by 66±5.0% (p<0.05) (Fig 2.7C; Fig 2.8). These data demonstrate that both the basic patch and the downstream QPV motif are involved in binding to importin β2. They suggest the NLS in N-terminus of anillin is a non-classical basic PY-NLS that contains a downstream (QPV) motif instead of the classical (PY) motif.
28 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.6
In vitro binding assays mapping where importin β2 binds to anillin. (A) Cartoon of the different anillin fragments fused to MBP used in this study. (B) Western-blotting analysis to identify co-purifying MBP-anillin fragments in a 6xHis-importin β2 pull-down assays. 6xHis-importin β2 was pulled down and co-purifying MBP-anillin fragments were analyzed.
29 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.7
In vitro binding assays between anillin mutants and importin β2. (A) Cartoon of anillin (1-151) fragments containing mutations and the wild type fused to MBP and analyzed for binding to importin β2. (B) (C) Western-blotting analysis showing co-purifying MBP-anillin (wild-type and mutants) in 6xHis-importin β2 pull-down assays. 6xHis-importin β2 was pulled down and co-purifying MBP-anillin was analyzed.
30 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.8
Quantitation of blot intensities of the pull down groups in Fig 2.7. Anillin mutants here are indicated by the numbers: (1)-KKR(68-70)-AA, (2)-PV(87-88)-AA, (3)-KKR(68-70)- AAA+PV(87-88)-AA, (4)-E83A, (5)-N84A, (6)-Q86A. Three parallel experiments were repeated for each group and blot intensities were compared to the wild-type binding group. The blot intensity of wild-type binding group was set to 1.0. Error bars indicate ± SEM.
31 Chapter 2 Identification of the anillin nuclear targeting sequence
2.2.6 Importin β2 targets anillin to the nucleus in interphase
To determine if importin β2 binding is required for the nuclear targeting of anillin in interphase, I generated full-length (FL) anillin and N-terminal 1-300 fragment of anillin with the mutation KKR (68-70)-AAA fused to GFP. Wild type and mutant constructs were transiently expressed in cells and their subcellular localization visualized by the GFP signal (Fig 2.9A). GFP-anillin with wild-type sequence localized to the nucleus (Fig 2.9B). In contrast, FL or the 1-300 containing the mutation KKR (68-70)-AAA failed to concentrate in the nucleus (Fig 2.9B). The difference in construct localization was further analyzed by comparing the fluorescence intensity of GFP signals of the nucleus to the cytoplasm (Fig 2.10). The mutants containing constructs localizing to cytoplasm rather than the nucleus had a significantly lower ratio of fluorescence intensity (nucleus:cytoplasm) than wild-type constructs that localized to the nucleus (Fig 2.10). These data demonstrate that importin β2 binding is required for the nuclear targeting of anillin to the nucleus during interphase. They indicate that the KKR (68-70)-x(15)-QPV (86-88) on N-terminal of anillin is the PY-NLS-like motif determines importin β2 binding.
32 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.9
Importin β2 targets anillin to the nucleus in interphase. (A) Cartoon of full-length or fragments of anillin with or without mutations at KKR (68-70) fused to GFP and expressed in cells. (B) Immunofluorescence images showing the localization of different constructs in interphase. Scale bars=10µm.
33 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.10
Quantitation of fluorescence intensities of different constructs. The ratio of fluorescence intensities between nucleus and cytoplasm were quantitated for each group. More than 100 cells were quantitated in each group (n=3). Error bars indicate ± SEM.
34 Chapter 2 Identification of the anillin nuclear targeting sequence
2.2.7 The anillin-importin β2 interaction has no significant role in cytokinesis
To determine if the nuclear targeting of anillin contributes to its cellular roles, I first used RNAi rescue experiments to examine whether importin β2 binding has any effect on the cytokinetic functions of anillin. I generated full-length anillin either containing the mutation KKR (68-70)-AAA or the wild-type and fused them to GFP. Generation of stable cell lines expressing GFP-anillin-FLwt and GFP-anillin-FL-KKR (68-70)-AAA from a Tet-inducible promoter was achieved using the Flp-In system (Invitrogen). The anillin siRNA is designed against the 3’UTR of the endogenous anillin transcript (a sequence not present in the transcript generated from the plasmids) and therefore the GFP-anillin fusions expressed from the plasmids are siRNA resistant (Fig 2.11) (Liu et al. 2012; Renshaw et al. 2014).
Cell lines expressing mutant and wild-type anillin were analyzed after siRNA rescue experiments. The lysates were first analyzed by SDS-PAGE and Western-blotting to test the level of anillin expression (Fig 2.12). In the control siRNA groups without doxycycline, only anillin (endogenous) was expressed (Fig 2.12). In the control siRNA groups with doxycycline, anillin (endogenous) and GFP-anillin (mutant or wild-type) were expressed at comparable level (Fig 2.12). In the anillin siRNA groups without doxycycline, no anillin was expressed (Fig 2.12). In the anillin siRNA groups with doxycycline induction, only GFP-anillin (mutant or wild-type) was expressed (Fig 2.12). These data demonstrate the specificity and effectiveness of the anillin siRNA. They also demonstrate that endogenous anillin expression could be replaced by the expression of GFP-anillin constructs at comparable level to endogenous anillin.
To characterize the localization of GFP-anillins in the cell lines, cells were treated with doxycycline to induce ectopic expression of GFP-anillin constructs. Both GFP-anillin-FLwt and GFP-anillin-KKR (68-70)-AAA were expressed (Fig 2.13A, left panel). Wild-type anillin localized to the nucleus, and the mutant anillin localized to the cytoplasm (Fig 2.13A, left panel). To determine if importin β2 binding to anillin had a role in cytokinesis, mutant or wild-type anillin were expressed in cells depleted of endogenous anillin using siRNA (Fig 2.13A,B). One of the typical consequences of cytokinetic failure is the generation of
35 Chapter 2 Identification of the anillin nuclear targeting sequence bi-nucleate or multi-nucleate cells (Liu et al. 2012). In control siRNA treated group, the levels of bi-nucleate cells in both mutant and wild-type groups were very low (5.0%±0.5 in mutant group and 4.0%±0.5 in wild-type group) (Fig 2.13A,B). In anillin siRNA treated groups without doxycycline (no anillin in the cell), there was a significant increase in bi-nucleate cells (78%±8.0 in mutant group and 81%±7.0 in wild-type group) (Fig 2.13A,B), which was consistent with previous result of anillin RNAi experiments (Liu et al. 2012). In contrast, in anillin siRNA treated groups with doxycycline (GFP-anillin transgenes expressed), the bi-nucleate phenotype was rescued (23%±4.0 in mutant group and 20%±5.0 in wild-type group) (Fig 2.13A,B). These data suggest that the anillin mutant lacking the NLS functions normally in cytokinesis. Therefore the anillin-importin β2 interaction has no significant role in cytokinesis.
Fig 2.11
Cartoon of RNAi rescue experiments in this study. Anillin siRNA (see chapter 5.7.3) was designed to specifically against the 3’UTR of the endogenous anillin transcript and left the GFP-anillin fusions unaffected.
36 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.12
Test of expression of anillin in RNAi rescue experiments. Western-blotting analysis of the expression level of endogenous anillin and GFP-anillin in different groups. The bands of GFP-anillin and endogenous anillin were indicated as arrows. GFP-wt=full-length wild-type anillin fused to GFP. GFP-KKR (68-70)-AAA=full-length anillin with mutations at KKR (68-70)-AAA fused to GFP.
37 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.13
Importin β2 binding to anillin is not required for successful cytokinesis. (A) Immunofluorescence images showing the localization of different GFP-anillin constructs in the presence and absence of endogenous anillin. (B) Quantitation of the percentage of bi-nucleate cells in each experimental group. More than 100 cells in each group were counted (n=3). Error bars indicate ± SEM. Scale bars=10µm.
38 Chapter 2 Identification of the anillin nuclear targeting sequence
2.2.8 The nuclear targeting of anillin contributes to maintain normal cell shape
Although the anillin-importin β2 interaction primarily regulates the nuclear targeting of anillin and has no role in cytokinesis, a basic question of why the cells have to enrich anillin in the nucleus is unknown. As discussed in chapter 1.2.5, one possible answer is to sequester anillin away from the interphase cytoplasm to prevent it disrupting cytoplasmic function. To determine if the nuclear targeting of anillin contributes to other cellular properties in interphase, full-length mutant anillin lacking the NLS and wild-type anillin were overexpressed in HeLa cells by transient transfection. Cells overexpressing wild-type anillin to the interphase nucleus had a normal round-cell shape (Fig 2.14). In contrast, cells overexpressing mutant anillin lacking the NLS which localized to cytoplasm had an abnormal shape; they appeared to be narrower in size with irregular protrusions (Fig 2.14). In addition, the fluorescence intensity of actin filaments seemed stronger in mutant expressing cells with abnormal cell shape, suggesting increasing levels of actin polymerization in these cells.
To compare the differences in cell shapes, the ratios of the width (‘a’ in Fig 2.15A) to the length (‘b’ in Fig 2.15A) of cells were measured. Wild-type anillin expressing cells and non-transfected cells have an a:b value close to 1.0 (Fig 2.15A). In contrast, mutant anillin expressing cells have a significantly lower a:b value compared to wild-type cells (Fig 2.15A). These data demonstrate excessive cytoplasmic anillin disrupts normal cell shape.
39 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.14
Overexpression of cytoplasmic anillin induces abnormal cell shape. Immunofluorescence images showing the localization of different GFP-anillin constructs and the actin cytoskeleton. Anln-FLwt=full-length wild-type anillin fused to GFP. Anln-FLmut=full-length anillin with mutations at KKR (68-70)-AAA fused to GFP. Scale bars=10µm. Cells expressing mutant anillin are indicated by arrows.
40 Chapter 2 Identification of the anillin nuclear targeting sequence
Fig 2.15
Quantitation of cell shape upon anillin expression. (A) Cartoon outlining the cell parameters measured to determine the ratio of cell width (a) to length (b). (B) Quantitation of the ratio of width (a) to length (b) for each group. More than 100 cells were quantitated in each group (n=3). Error bars indicate ± SEM.
41 Chapter 2 Identification of the anillin nuclear targeting sequence
2.3 Discussion
2.3.1 The non-typical PY-NLS in Nab2 and anillin
Human importin β2 recognizes both the hydrophobic and basic subclasses of PY-NLSs, while the yeast homolog importin 104p recognizes only basic PY-NLSs (Chook and Suel 2010). The basic residues upstream of the (PY) motif were required for importin β2 binding (Cansizoglu et al. 2007; Suel et al. 2008). Mutating arginine residues to alanines in the upstream basic patch of the Hrp1p PY-NLS significantly reduced importin β2 binding by 5-fold (Suel et al. 2008). In addition, previous studies have revealed a yeast protein Nab2, that contains a (PL) instead of a typical (PY) motif as part of the PY-NLS (Suel et al. 2008; Soniat et al. 2013). Crystallography studies of the importin β2-Nab2PY-NLS complex have shown that Pro-238 of Nab2 interacts primarily through hydrophobic interactions with sidechain residues of importin β2 (Soniat et al. 2013). Leu-239 of Nab2 makes hydrophobic interactions with importin β2 (Soniat et al. 2013). Phe-236, which resides in the N-terminal of the (PL) residues, also makes hydrophobic interaction with importin β2 (Soniat et al. 2013). In addition, the neighboring Arg-235 makes multiple salt bridges and hydrogen bonds with importin β2. These results showed in Nab2 PY-NLS the typical (-PY-) motif can be replaced by (-Pϕ-) where ϕ is a hydrophobic residue that contributes to bind to importin β2 (Suel et al. 2008; Soniat et al. 2013). Meanwhile, multiple neighboring residues of the (-Pϕ-) also contribute to the interaction to importin β2 (Soniat et al. 2013). In the case of anillin, a (QPV) motif instead of the typical (PY) contributes to importin β2 binding. One possible model is that Val-88 makes hydrophobic interaction with importin β2 in addition to the interaction between Pro-87 and importin β2. Moreover, Phe-82 that resides in the N-terminal of the (QPV) motif may also contribute to importin β2 binding in a similar mechanism as Phe-235 of Nab2 does.
I identified a novel non-classical NLS in anillin and demonstrated that anillin binds to importin β2 through a variant of the basic-PY motif that contains a (PV) instead. PY-NLSs were suggested to vary in residues and structures in individual cases (Lee et al. 2006; Cansizoglu et al. 2007; Suel et al. 2008). To further compare the PY-NLS like motif of anillin
42 Chapter 2 Identification of the anillin nuclear targeting sequence to other non-typical PY-NLS, I have listed three non-typical PY-NLS and aligned them with other similar typical PY-NLS (Fig 2.16). It is interesting to note that Valine, Leucine and Glutamine are always next to the core (Pϕ) motif where ϕ is hydrophobic residue (Fig 2.16). My data broaden our knowledge of PY-NLS motif and the molecular mechanism of importin β2-cargo interaction. My data support a model whereby the (PY) motif is not absolutely required for importin β2 binding. My data also highlights that there is greater potential variation in PY-NLS than previously thought and therefore a new search will be needed to identify all importin β2 binding substrates rather than the previous search for the putative (PY) motif.
Fig 2.16
Alignment of PY-NLS sequences. Alignment of the 3 (Pϕ)-NLS with 3 PY-NLSs that have similar neighboring residues to the core (-Pϕ-) motif. The upstream basic patches and the downstream (PY) residue are marked in red. Residues that neighbor the proline or the (PY) motif are marked in purple.
43 Chapter 2 Identification of the anillin nuclear targeting sequence
2.3.2 The abnormal cell shape caused by cytoplasmic anillin
In this study, the biological meaning of the nuclear targeting of anillin was further revealed. Anillin targeting to the nucleus prevented anillin from disrupting cytoplasmic processes that regulate cell shape, like actin cytoskeleton organization. Cells overexpressing excessive cytoplasmic anillin had abnormal cell shape and stronger fluorescence intensity of actin filaments, suggesting excessive cytoplasmic anillin may induce hyper-polymerization of actin (Fig 2.14). However, the mechanism of how cytoplasmic anillin alters the global actin filament organization needs to be further analyzed. Other cellular factors that are involved in modulating actin filament organization should be considered. mDia2 and its activator RhoA are great candidates since mDia2 is an actin nucleator downstream of RhoA (Chapter 4.2.4a) and at the same time mDia2 and RhoA both bind to anillin (Piekny et al. 2008; Watanabe et al 2010; Liu et al. 2012). I’ve tried to analyze the localization of RhoA and mDia2 in cells overexpressing cytoplasmic anillin but the initial experiment was unsuccessful. In future studies I will examine the actin hyper-polymerization phenotype in cells overexpressing cytoplasmic anillin and determine if it is dependent on RhoA and mDia2. I will inhibit the activity of RhoA using Clostridium botulinum C3 exoenzyme or using mDia2 RNAi to deplete mDia2 in mutant cells. Then I will identify if and how RhoA and mDia2 are involved in stimulating actin polymerization in cells overexpressing cytoplasmic anillin.
In addition, the thinner and more irregular shape of the cells expressing mutant anillin suggests the excessive cytoplasmic anillin breaks the balance of global cytoskeletal tensions. One possibility is that excessive cytoplasmic anillin disrupts the distribution and function of anillin binding partners such as the septin cytoskeleton (Kinoshita et al. 2002; Liu et al. 2012). I’ve tried to analyze the septin localization but the initial experiment was unsuccessful. Next I’ll optimize fixation and staining conditions to determine if the septin cytoskeleton is disrupted in mutant-expressing cells. These experiments will reveal how excessive cytoplasmic anillin affects the factors that regulate cell shape in interphase. More importantly, it may reveal a broadly applicable mechanism of how cells utilize a nuclear targeting mechanism to prevent proteins from disrupting certain cellular process in interphase.
44 Chapter 3 The role of anillin in cilia function
Chapter 3. The role of anillin in cilia function
3.1 Introduction
Motile cilia move fluids and cells, and nonmotile cilia are involved in receiving and transducing signals for cell development and homeostasis (Baker and Beales 2009; Goetz and Anderson 2010). In vertebrate cells, cilia are antenna-like structures projecting from the cell surface and arise from a basal body that is anchored to the base of the ciliary membrane by transition fibers (Reiter et al. 2012). Cilia are compartmentalized from the rest of the cell by a complex structure consisting of transition fibers and transition zones that link the proximal part of the axoneme (microtubule-based skeleton of cilia) to the ciliary membrane (Reiter et al. 2012). Protein transport into cilia is regulated at the ciliary base (Nachury et al. 2010). Septins, which are anillin binding partners (Kinoshita et al. 2002; Liu et al. 2012), were initially implicated in forming a ‘diffusion barrier’ at the ciliary base that blocks free exchange between cilia and the rest of the cell (Hu et al. 2010). However a more recent study disputed this idea by showing localization of septin 2, 7 and 9 to the axenome instead of the basal body (Ghossoub et al. 2013). There was also evidence suggesting components of transition zone were involved in establishing the diffusion barrier (Chih et al. 2012).
The transport of cytosolic cargos into cilia is mediated by a size-selective diffusion barrier at the ciliary base (Kee et al. 2012). Inside the ciliary compartment, cargos are actively transported by the intraflagella transport (IFT) machinery and ciliary motors such as kinesins and dyneins (Deane et al. 2001; Qin et al. 2005; Hao et al. 2011). Studies have suggested similarities between the ciliary and nucleocytoplasmic transport machineries (Hurd et al. 2011; Fan et al. 2011; Dishinger et al. 2010). Importins (β1 and β2) were reported to be involved in cargo transport to the cilia, coupled with the RanGTP gradient across the basal body (Hurd et al. 2011; Dishinger et al. 2010; Fan et al. 2007). However, a more recent study has suggested the diffusion barrier is mechanistically distinct from those of the nuclear pore complex (NPC) (Breslow et al. 2013). Therefore, if and how the Ran-NTR mechanism functions in cilia is unclear. A previous study reported that importin β2 targeted the kinesin-2 KIF 17 to cilia through a cilia localization sequence (CLS) (Dishinger et al. 2010). The CLS
45 Chapter 3 The role of anillin in cilia function of KIF17 contains an upstream basic patch of residues with a downstream (PL) motif that is recognized by importin β2, instead of the classical (PY) motif (Dishinger et al. 2010). The CLS of KIF17 bares resemblance to the PY-NLS like motif of Nab2 (yeast) and anillin (human); they all contain (PΦ) motif instead of (PY) where Φ are hydrophobic residues (Fig 3.1) with upstream basic patches of residues and that bind to importin β2 (Dishinger et al. 2010; Soniat et al. 2013; Chapter 2.2.5). Based upon current literature and in light of my discovery that anillin, like KIF17, has an importin β2 binding site, I hypothesized that anillin could be localized to the cilia.
Fig 3.1
Sequence alignments of non-PY motifs of anillin, Nab2, KIF17. The upstream basic patches and the downstream proline residue are marked in red. Residues that neighbor the proline and contribute to importin β2 binding are marked in purple.
46 Chapter 3 The role of anillin in cilia function
3.2 Results
3.2.1 Anillin localizes to the basal body of the cilia
To determine if anillin is targeted to cilia, I examined the localization of anillin in hTERT RPE-1 cells that were serum starved to induce differentiation and cilia formation. Cells were fixed and analyzed by IF and confocal microscopy using antibodies against acetylated tubulin (to identify cilia), γ-tubulin antibody (to identify the cilia basal body), and anillin. In differentiated cells, anillin localized to the basal body of the cilia (Fig 3.2). In very rare cases anillin was observed as dots along the antenna-like cilia structure (Fig 3.2). These data suggest anillin may reside in the cilia, but mostly localized to the basal body and in very rare cases localized along the antenne-like cilia structure.
3.2.2 Anillin RNAi cause defect in cilia length
To determine if anillin has a role in ciliogenesis or maintenance of cilia I analyzed the effect of anillin depletion on cilia. An anillin RNAi experiment was carried out using differentiated hTERT RPE-1 cells. Cells were treated with anillin RNAi and lysates were generated and analyzed by SDS-PAGE and Western-blotting to test the level of expression of anillin. In anillin RNAi treated group, there was no detectable expression of anillin (Fig 3.3A). In the control group of cells, the expression of anillin was not affected (Fig 3.3A). These data demonstrate anillin was depleted in differentiated hTERT RPE-1 cells treated with anillin RNAi.
To confirm the anillin staining at basal bodies was dependent on the expression of anillin, antibody staining was carried out on differentiated hTERT RPE cells treated with control or anillin siRNA. In control siRNA treated cells, anillin localized to the basal body, a localization consistent with my observation in untreated cells (Fig 3.3A). In contrast, in anillin siRNA treated cells, anillin was absent in both the nucleus and the basal body (Fig 3.3B). Moreover, in anillin siRNA treated group, cilia were shorter than those in control group (Fig 3.3B). These data suggest anillin may have a role in ciliogenesis or maintenance
47 Chapter 3 The role of anillin in cilia function of cilia.
Fig 3.2
Anillin localizes to cilia in differentiated hTERT RPE-1 cells. Immunofluorescence of serum starved hTERT RPE-1 cells by confocal microscopy showing the localization of anillin. Scale bars=10µm.
48 Chapter 3 The role of anillin in cilia function
Fig 3.3
Anillin RNAi in differentiated hTERT RPE-1 cells. (A) Western-blotting analysis showing the level of expression in each group. (B) Immunofluorescence of differentiated hTERT RPE-1 cells by confocal microscopy showing the localization of anillin in control siRNA (top panel) and the anillin siRNA treated group (bottom panel). Scale bars=10µm.
49 Chapter 3 The role of anillin in cilia function
3.2.3 The localization of anillin changes during cell differentiation
Interestingly, in differentiated cells where anillin localized to the basal body of cilia, there was no nuclear localization of anillin (Fig 3.2). In undifferentiated cells, anillin localized to the nucleus (Fig 2.1; Fig 3.2). To confirm anillin is targeted to the nucleus in undifferentiated cells, hTERT RPE-1 in the presence of serum were fixed and stained with an anti-γ-tubulin antibody and an anti-anillin antibody. Cells were then analyzed by confocal microscopy. In contrast to differentiated cells where anillin localized to the cilia (Fig 3.4 bottom panel), anillin localized to the nucleus in differentiated cells (Fig 3.4 top panel). These results demonstrate the change in localization of anillin during cell differentiation; anillin localizes to nucleus in undifferentiated cells but not in differentiated cells where it instead localizes to the basal body of cilia.
These results raise important questions about why anillin localizes differently during differentiation and how it is regulated. I hypothesized several possibilities:
(1). Signaling or post-translational modification prevents anillin being targeted to the nucleus.
(2). Differentiation causes the expression of alternative isoforms of anillin that lack an NLS.
(3). Importin β2 is differentially regulated during differentiation; switching from nuclear transport to ciliary transport thereby targeting anillin to the cilia.
(4). The nuclear targeting and ciliary targeting of anillin is regulated by different receptors.
50 Chapter 3 The role of anillin in cilia function
Fig 3.4
The localization of anillin is regulated during differentiation. (A) Immunofluorescence images analyzed by confocal microscopy showing the localization of anillin in undifferentiated (top panel) and differentiated (bottom panel) hTERT RPE-1 cells. Scale bars=10µm.
51 Chapter 3 The role of anillin in cilia function
3.2.4 Anillin may not be an alternative isoform in differentiated cells
Anillin is absent from the interphase nucleus in differentiated cells (Fig 3.4). To determine if an alternatively spliced isoform of anillin lacking the NLS is expressed during differentiation, undifferentiated and differentiated hTERT RPE-1 cells were harvested and analyzed for the expression of anillin. The molecular weight and the level of expression of anillin did not change during differentiation (Fig 3.5). These results suggest that the anillin at the cilia basal body is the same or at least very similar to anillin in undifferentiated cells that has the NLS.
3.2.5 Phospho-mimic mutation does not affect the anillin-importin β2 interaction
My result above suggests that anillin targeting to the cilia instead of nucleus is regulated by a different mechanism rather than alternative splicing of anillin. An alternative hypothesis is that the NLS of anillin is switched off in differentiated cells. It has been reported that the conserved serine residue at 72 (S72) that lies within the mapped NLS is phosphorylated in mitosis (Fig 2.3) (Dephoure et al. 2010). I hypothesized the phosphorylation of S72 could regulate anillin-importin β2 binding by introducing a negative charged residue very close to a basic patch of residues that are required for the anillin-importin β2 binding (Chapter 2.2.5). To determine whether phosphorylation on S72 regulates the the anillin-importin β2 interaction, I mutated S72 of anillin to alanine or aspartate (to mimic the negative charge of a phosphate) and analyzed the anillin mutant binding to importin β2 (Fig 3.6). Anillin fragment 1-151 containing the S72D or the S72A mutation did not reduce the binding to importin β2 when compared to the wild-type anillin fragment 1-151 (Fig 3.6). In contrast, an anillin fragment lacking the first 1-91aa reduced binding to importin β2 (Fig 3.6). These data suggest that phosphorylation on S72 may not regulate the anillin-importin β2 interaction. Therefore another mechanism is likely involved in regulating the localization of anillin during cell differentiation.
52 Chapter 3 The role of anillin in cilia function
Fig 3.5
The molecular weight and level of expression of anillin do not change during cell differentiation. Western-blotting analysis showing the molecular weight and level of expression of anillin in differentiated and undifferentiated cells.
Fig 3.6
In vitro binding assays between anillin phospho-mimic mutant and importin β2. Western-blotting analysis showing co-purifying MBP-anillin (wild-type and mutants) by 6xHis-importin β2 pull-down assays. 6xHis-importin β2 was pulled down and co-purifying MBP-anillin was analyzed. The bands of anillin are indicated by arrows.
53 Chapter 3 The role of anillin in cilia function
3.3 Discussion
In this study, I discovered the ciliary localization of anilin, which has not been identified before. Anillin localizes to the basal body of cilia, instead of nucleus, in differentiated cells. To identify the ciliary targeting mechanism of anillin, I first investigated if anillin is differentially regulated during differentiation. One possibility is differentiation causes the expression of alternative isoforms of anillin that lack the NLS, however my data suggest anillin is not truncated in differentiated cells. Another possibility is the NLS of anillin is switched off during differentiation. I investigated it by generating a phospho-mimic mutation at S72 within the NLS and identified it did not effect importin β2 binding. My data does not exclude a model whereby importin β2 by itself may be regulated during differentiation either in its level of expression or its association with different subcellular structures (nuclear pore complex or the basal body complex). Therefore importin β2 could be functionally switched from nuclear transport to cilia transport and targets anillin to the cilia. Understanding this ciliary targeting mechanism will reveal how a cell uses two different transport pathways during differentiation. In addition, studies have suggested NTR-RanGTP/GDP pathway was involved in ciliary transport while others disputed it (Fan et al. 2007; Hurd et al. 2011; Breslow et al. 2013). Identifying the role of the anillin-importin β2 interaction in ciliary targeting of anillin will further reveal how cargos are transported to the cilia. It will also reveal the relationship between the nuclear transport and ciliary transport.
In the next step, I will test if the level of importin β2 is regulated and identify the localization of importin β2 during cell differentiation to test if importin β2 is differentially regulated during differentiaton. Secondly, I will analyze the localization of the NLS mutant of anillin during cell differentiation. If the mutant does not localize to cilia, it indicates anillin and importin β2 plays a role in ciliary targeting of anillin. However if the mutant localizes to the cilia but not to the nucleus, it indicates another receptor could be involved in targeting anillin to the cilia.
It is interesting to identify the presence of anillin to the basal body of cilia as it will broaden our knowledge about cilia and ciliogenesis. Anillin depletion in hTERT RPE-1 cells resulted
54 Chapter 3 The role of anillin in cilia function in shorter cilia, suggesting a role of anillin in cilia morphology. Studies have shown the involvement of septins during ciliogenesis, although there is dispute as to when and where they contribute to this process (Hu et al. 2010; Ghossoub et al. 2013). The detailed role of anillin in cilia cannot be inferred at this time. One possibility is that, since anillin binds to septins (Kinoshita et al. 2002; Liu et al. 2012), it has a relationship with the septin function in cilia. To further identify the roles of anillin in ciliogenesis, I will use time-lapse imaging to determine the changes of the localization of anillin and anillin binding partners such as septin and importin β2 during ciliogenesis process. It will reveal the role of anillin in ciliogenesis in time and space.
55 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Chapter 4. Anillin enhances RhoA dependent activation of mDia2
4.1 Introduction
4.1.1 Formins and actin nucleation
Dynamic cellular events including shape changes, polarization, adhesion, migration, and cell division depend on the remodeling of the actin cytoskeleton (Campellone and Welch 2010). Consequently, actin polymer formation is tightly regulated in space and time (Chesarone et al. 2010). Actin nucleators that can actively catalyze actin filament assembly play pivotal roles in directing cellular actin cytoskeleton organization. Formins are one of the three bona fide actin nucleators (Formins, Arp2/3 and Spire) and nucleate the formation of linear unbranched actin filaments (Pruyne et al. 2002; Sagot et al. 2002). The nucleation mechanisms of three classes of nucleators vary greatly. In general, formins stabilize actin polymerization intermediates (dimers), and Arp2/3-WASp-actin complex can mimic an actin trimer, while Spire organizes actin monomers into a pre-nucleated template (Fig 4.1) (Goode and Eck 2007).
The Diaphanous formins (mDia1, mDia2, mDia3) belong to the diaphanous-related formins (DRF) family that includes most functional formins in metazoans. Typically a DRF molecule is divided into the C-terminal region containing the FH2 domain that nucleates actin filaments, and the N-terminal regulatory region (Fig 4.2). In vivo and in vitro studies have shown that the FH2 domain directly nucleates actin filaments (Evangelista et al. 2002, Sagot et al. 2002). The formin FH2 domain forms a unique flexible tethered dimer and interacts with the barbed end to nucleate the filaments with a processive capping effect in a “stair-step” way (Xu et al. 2004; Moseley et al. 2004; Pruyne et al. 2002). The FH1 domain binds to profilin and helps FH2 domain to facilitate the profilin-actin monomer polymerization and increases the rate of filament elongation (Pollard et al. 2000). The profilin-actin complex is the predominant form of actin monomer for polymerization in most eukaryotic cells (Pollard
56 Chapter 4 Anillin enhances RhoA dependent activation of mDia2 et al. 2000). Studies have shown FH1-FH2 domains together can utilize monomeric profilin-actin for polymerization. Disrupting the profilin-FH1 interaction reduces the efficiency of actin assembly (Sagot et al. 2002; Pring et al. 2003).
Fig 4.1
Three different nucleation mechanisms. In spontaneous actin assembly, the actin dimers and trimers are unstable and rapidly dissociate. The dimer/trimer will only proceed to polymerize further at high actin monomer concentrations. Formins stabilize actin polymerization intermediates and nucleate linear unbranched actin filaments. The Arp2/3 complex associated with WASp-actin can mimic an actin trimer and nucleates the formation of a new filament from an existing actin filament. Spire recruits and organizes actin monomers into a stable pre-nucleated template. (Figure adapted with modification from Goode and Eck 2007)
57 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.2
Formin domain structure and activation mechanism. The C-terminal FH2 domain of formins is the actin nucleation site. The C-terminal FH1 domain can interact with profilin-actin and induce filament elongation. The N-terminal GBD domain binds to RhoGTP and promotes the release of the autoinhibitory binding between the N-terminal DID and the C-terminal DAD domains, leading to the activation of formins. (Figure adapted with modification from Goode and Eck 2007)
58 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
4.1.2 Regulation of formin activation
The activity of formins has to be regulated to dictate when and where actin polymer is formed and to keep the balance between the assembly and disassembly of cellular actin polymers. However, the detailed mechanism of formin activation remains unclear in metazoan cells. Diaphanous formins are auto-inhibited by an interaction between the diaphanous inhibitory domain (DID) in the N-terminal region and the diaphanous autoregulatory domain (DAD) in the C-terminal actin nucleation region (Fig 4.2) (Li and Higgs 2003). Small G-proteins (like Rho) can bind to the G-protein binding domain (GBD) that neighbors the DID domain and this is proposed to trigger the release of the DID-DAD auto-inhibitory binding (Li and Higgs 2005; Lammers et al. 2005). Crystallography studies show that Rho binding induces a conformational change in GBD and DID domains that occlude the DAD-DID interaction surface (Lammers et al. 2005). The residues of Rho also interfere and repulse the DAD from the DID domain (Lammers et al. 2005; Nezami et al. 2006).
However, biochemical studies show that Rho binding on its own is not sufficient to fully activate the actin nucleation activity of mDia2. In vitro studies find that, either using fragments of mDia1 or full-length mDia1, the activation effect of Rho is only partial even when a thousand fold molar excess of Rho is used (Li and Higgs 2005; Maiti et al. 2012). In addition, an in vivo study has shown that, the function of RhoA-mDia1 signaling pathway in microtubule stabilization in mouse fibroblasts was facilitated by integrin-mediated activation of focal adhesion kinase (FAK) at the leading edge (Palazzo et al. 2004). These results suggest there are factors in addition to Rho that are involved in activating formins under physiological conditions. During animal cell cytokinesis, formins are recruited to the furrow and co-localize with Rho (Watanabe et al. 2008). Therefore the additional activators may be expected to localize at the ring structure to cooperate with Rho in activating formins.
59 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
4.1.3 Formins in cytokinesis
4.1.3.1 The pivotal roles of formins in cytokinesis
The dynamic remodeling of the actin cytoskeleton is essential for actomyosin ring organization and furrow ingression during cytokinesis. Studies in S.pombe and Drosophila cells revealed the pivotal roles for formins during actomyosin ring assembly, stabilization and constriction (Wu and Pollard 2005; Castrillon and Wasserman 1994). Moreover, electron microscopic analysis finds that linear actin filaments are the main component of the ring, which suggests the participation of formins in the ring formation as formins generate linear filaments (Maupin and Pollard 1986; Sagot et al. 2002).
4.1.3.2 The potential role of anillin in mDia2 activation
In human cells, mDia2 is indispensible for maintaining correct furrow position and stabilizing the ingression (Watanabe et al. 2008). mDia2 is recruited to the membrane furrow where RhoA is enriched (Watanabe et al. 2010; Hickson and O’ Farrell 2008). Both RhoA and anillin are required for the targeting of mDia2 to the furrow, yet how the activation of mDia2 is regulated to coordinate with cytokinetic ring organization is unclear. Anillin binds to the DID domain and prevents the binding of the DAD domain to the DID domain (Watanabe et al 2010). Based on these studies, I propose a novel model where anillin is an activator of mDia2. Anillin could operate alone or as a co-activator/enhancer in conjunction with RhoA. Since the presence of RhoA at the furrow is not sufficient to activate mDia2, it is reasonable to propose anillin anchors mDia2 within the ring structure while at the same time working in association with RhoA to provide localized activation for mDia2.
4.1.4 Objectives and strategies
It is a well-established fact that anillin acts as an actomyosin filament binding protein during cytokinesis (reviewed in Piekny and Maddox 2010). However, the idea that anillin could also be a regulator of the cellular actin cytoskeleton via its potential activating effect on mDia2 is exciting. First I will characterize the anillin-mDia2 interaction and its effect on the DAD-DID
60 Chapter 4 Anillin enhances RhoA dependent activation of mDia2 autoinhibitory binding. Next I will test if anillin has a role in activating mDia2 using in vitro actin polymerization assays. The ultimate goal is to identify how anillin-mDia2 interaction contributes to cytokinesis and reveal novel regulatory mechanisms of the actin cytoskeleton.
61 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
4.2 Results
4.2.1 mDia2 has a distinct binding site within region N2 of anillin
Previously the mDia2 binding site was defined as residing in the first 91 amino acids of anillin (Watanabe et al. 2010). This region of anillin also contains the CD2AP binding site in region N1 (Monzo et al. 2005) and the importin β2 binding motif in N3 and N4 that I discovered and described earlier (Fig 4.3) (Chapter 2). To identify where mDia2 binds I generated anillin fragments 1-57, 1-91, 1-151, 1-300, 57-91, 57-151, 22-151, fused to MBP and analyzed their binding to the mDia2 fragment NT89-533, fused to GST (Fig 4.4A). Proteins were incubated, then re-isolated on glutathione beads and analyzed by Western-blotting using an anti-MBP antibody to detect co-purifying anillin fragments. Anillin fragments containing amino acid residues 22-91 bound to mDia2, while fragments lacking 22-91 did not (Fig 4.4B). These data suggest that amino acids 22-91 of anillin form the predominant site for mDia2 binding.
To determine if mDia2 has a distinct binding site from those of CD2AP and importin β2 that also binds to the 1-91aa of anillin, the mDia2 fragment NT89-533 fused to GST was incubated with the anillin fragment 1-151 fused to MBP in the presence of GST-CD2AP and 6xHis-importin β2. mDia2 and anillin were mixed first and then incubated with CD2AP or importin β2. Proteins were re-isolated on amylose resin and analyzed by Western-blotting using an anti-GST antibody to detect co-purifying GST-NT89-533. The amount of mDia2 co-purifying with anillin was not reduced in the presence of increasing concentrations of CD2AP or importin β2. The amount of GST-CD2AP and 6xHis-importin β2 loaded was up to 1-fold molar ratio of loaded mDia2 (Fig 4.5). These data suggest that CD2AP and importin β2 did not compete with mDia2 to bind to anillin. As the mDia2 binding site includes the N2 region and a C-terminal portion of N1 region (Fig 4.4B), while CD2AP binds to N1 region and importin β2 binds to N3-N4 regions, these data suggest that mDia2 has a distinct binding site within N2 region from amino acid residues 37-50 (Fig 4.3, marked with arrow).
62 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.3
Sequence alignment of first 100 amino acids of anillin. The potential mDia2 binding site is indicated by the arrow. The CD2AP and importin β2 binding sites are marked in triangle and asterisk, respectively.
63 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.4
In vitro binding assays mapping the mDia2 binding site on anillin. (A) Western-blotting analysis showing co-purifying MBP-anillin fragments by GST-NT89-533 pull-down assays.
GST-NT89-533 was pulled down and co-purifying MBP-anillin fragments were analyzed. (B)
Loading control of GST-NT89-533 and MBP-anillin fragments. The bands of anillin fragments (57-91), (57-151), (22-151) are indicated with arrows.
64 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.5
In vitro competition assays between importin β2, CD2AP and mDia2 in binding to anillin. (A) Western-blotting analysis showing co-purifying GST-NT89-533 by MBP/MBP- anillin pull-down assays in the presence of increasing concentrations of 6xHis-importin β2.
(Anln=anillin) (B) Western-blotting analysis showing co-purifying GST-NT89-533 by MBP/ MBP-anillin pull-down assays in the presence of increasing concentrations of CD2AP. (N-CD2AP=N-terminus of CD2AP including the first 3 SH3 domains)
65 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
4.2.2 Anillin enhances RhoA dependent inhibition of the mDia2 DID-DAD domain interaction mDia2 functions to stabilize furrow formation and ingression (Watanabe et al. 2008). Anillin is required for maintaining the furrow localization of mDia2 (Watanabe et al. 2010). Yet the mechanism of how mDia2 is activated at the furrow remains unclear. Most analysis of DRFs has been carried out on mDia1. RhoA binding to RBD domain of mDia1 only partially relieves the autoinhibitory DID-DAD interaction of mDia1 (Li and Higgs 2005; Maiti et al 2012). Therefore additional factors could be required to fully activate mDia1 (Li and Higgs 2005; Maiti et al. 2012). Interestingly, anillin was reported to bind to the DID domain and prevent the binding of DAD domain to DID domain of mDia2 (Wanatabe et al. 2010). Therefore, anillin could be involved in modulating mDia2 activity at the membrane furrow. I hypothesized three potential models of how anillin functions in relation to mDia2 (Fig 4.6). In the first model anillin recruits mDia2 but has no effect on mDia2 activation even with the presence of RhoA, a model based on the study of Watanabe et al. 2010 (Fig 4.6B). In the second model, anillin on its own can activate mDia2 (Fig 4.6C). In the third model, anillin by itself has no activation effect on mDia2, but works in conjunction with RhoA to co-activate/enhance mDia2 activation (Fig 4.6D).
To further test these models of the potential activation effects of anillin (and RhoA) on mDia2, I developed a competition assay to test the effect of anillin and RhoA on relieving mDia2 autoinhibition. First, I determined if mDia2 NT1-533 (N-terminal 1-533aa of mDia2) bound to CT (C-terminus of mDia2) in the same manner as mDia1. Then I tested the effect of
RhoA and anillin on the NT1-533-CT interaction (Fig 4.7). NT1-533 bound to CT and the addition of increasing concentrations of GST (up to 1.0 molar ratio to loaded CT) did not disrupt this interaction (Fig 4.8A). In contrast, increasing concentrations of GST-RhoA (up to
1.0 molar ratio to loaded CT) partially disrupted the interaction of NT1-533 and CT (Fig 4.8A). However, increasing concentration of 6xHis-anillin (up to 1.0 molar ratio to loaded CT) had no significant inhibitory effect on the interaction of NT1-533 and CT (Fig 4.8A). Interestingly, addition of anillin in the presence of RhoA enhanced the inhibitory effect of RhoA on the
NT1-533-CT interaction (Fig 4.8A). These data suggest that anillin by itself has no inhibition effect. Instead, anillin enhances the inhibitory effect of RhoA on the mDia2 DID-DAD domain interaction.
66 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.6
Cartoon of the proposed models for the potential roles of anillin in mDia2 activation. (A) Anillin concentrates at the membrane furrow and recruits mDia2. The autoinhibitory binding between the DID-DAD domains keeps mDia2 in auto-inhibited state before activation. (B) In the recruiter model, anillin simply acts as a recruiter and has no role in activating mDia2. (C) In the activator model, anillin can recruit and activate mDia2 on its own. (D) In the enhancer model, anillin has no activation effect on mDia2, instead it works in conjunction with RhoA to activate mDia2.
Fig 4.7
Cartoon of the in vitro competition assays. The MBP-NT1-533 and GST-CT binding was analyzed in the presence of anillin and RhoA.
67 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.8
In vitro competition assays testing the effect of anillin and RhoA on the interaction between mDia2 fragments NT1-533 and CT. (A) Western-blotting analysis showing co-purifying GST-CT by MBP-NT1-533 pull-down assays in the presence of GST, GST- RhoA and 6xHis-anillin (1-300). Blot intensities in different groups were quantitated and compared to the wild-type pull-down group in the bottom panel. Each individual group was repeated three times. Error bars indicate ± SEM. (B) Loading control for MBP-NT1-533, GST-CT, 6xHis-anillin (1-300), GST, GST-RhoA and 6xHis-anillin (1-300).
68 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
4.2.3 Anillin does not bind to the NT1-533-CT complex
The biochemical results (4.2.2) demonstrate that anillin by itself does not release CT from
NT1-533, but enhances RhoA induced release of CT from NT1-533. However, the question of how anillin, RhoA and the NT1-533-CT complex interact with each other in time and space remains unclear. To determine if anillin binds to the NT1-533-CT complex, I first confirmed that anillin bound to NT1-533 but not the CT (Fig 4.9). Next, anillin was added to the pre-formed NT1-533-CT complex and analyzed for its ability to bind to the NT1-533-CT complex. Anillin was not pulled down by NT1-533 (Fig 4.10A), indicating it did not bind to the
NT1-533 in the presence of CT. In addition, RhoA was added to the pre-formed NT1-533-CT complex and analyzed for its binding to the NT1-533-CT complex. A small amount of RhoA was pulled down by NT1-533 (Fig 4.10B), indicating it bound weakly to NT1-533 and weakly displaced CT from the complex. In contrast, when anillin and RhoA were added together to the pre-formed NT1-533-CT complex, a greater amount of anillin and RhoA were pulled down by NT1-533 and displaced CT from the complex.
These data demonstrate that anillin does not bind to the NT1-533-CT complex, and RhoA only weakly interacts with the NT1-533 in the presence of CT. However, RhoA binding facilitates anillin binding to NT1-533 and the subsequent displacement of CT from the complex.
69 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.9
In vitro binding assays between anillin and NT1-533 and CT. (A) Western-blotting analysis showing co-purifying 6xHis-anillin by MBP/MBP-NT1-533 pull-down or GST/GST-CT pull-down assays. MBP/MBP-NT1-533 or GST/GST-CT was pulled down and co-purifying
6xHis-anillin was analyzed. (B) Loading control of MBP, MBP-NT1-533, GST and GST-CT.
The band of MBP-NT1-533 is indicated by the arrow.
70 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.10
Anillin binding to the DID domain is dependent on RhoA. (A) (B) Western-blotting analysis showing co-purifying 6xHis-anillin (or GST-RhoA) by MBP-NT1-533 that was co-purified by GST-CT pull-down assays (Anln=anillin, NT=NT1-533). GST-CT was pulled down and co-purifying MBP-NT1-533 and 6xHis-anillin (or GST-RhoA) were analyzed. (C)
Loading control of MBP-NT1-533, GST-CT, 6xHis-anillin and GST-RhoA-GTP. Bands in different pull-down groups are indicated by arrows.
71 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
4.2.4 Anillin enhances RhoA activation of mDia2 actin polymerization activity
The data in chapter 4.2.3 are highly suggestive of RhoA and anillin combining to modulate mDia2 activity. To assess the relative roles of RhoA and anillin in regulating mDia2 actin polymerization activity, I used a pyrene actin polymerization assay to analyze the extent of mDia2 activation in the presence of either RhoA or anillin (Kouyama and Mihashi 1981). Pyrene-labeled monomer actin is weakly fluorescent, but upon polymerization into actin filaments the fluorescence is enhanced up to 20 times (Fig 4.11A) (Tobacman and Korn 1983; Cooper et al. 1983). Therefore the polymerization process can be monitored in a fluorimeter. Typically for the in vitro actin polymerization assays, actin has a stable but very low self-polymerization rate. I confirmed that, addition of an actin nucleator (like gelsolin) greatly increases the polymerization rate (Fig 4.11B), as previously described (Arora and McCulloch 1996).
72 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.11
An example of the in vitro pyrene- actin polymerization assay. (A) Cartoon of the mechanim of the pyrene-actin polymerization assay. (B) An example of the in vitro pyrene-actin polymerization assay showing the self-polymerization of the actin and the polymerization reaction with the presence of gelsolin.
73 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
4.2.4.1 NT(1-533) inhibits the actin polymerization activity of CT
Previous studies demonstrated that mDia1 was able to induce actin filament formation in vitro (Li and Higgs 2005). To determine if mDia2 acts the same as mDia1, an in vitro pyrene-actin polymerization assay was carried out using the mDia2 fragment, CT. CT by itself induced actin polymerization (Fig 4.12A,C). In contrast, in the presence of equal molar
NT1-533, the actin polymerization activity of CT was inhibited (Fig 4.12B,C). The control proteins (GST or MBP) did not affect the actin polymerization activity of CT (Fig 4.12A,C). Therefore mDia2 behaves in the same way as mDia1; the CT that contains the FH2 domain is able to stimulate actin polymerization and this activity is inhibited in the presence of the NT that contains the DID domain.
4.2.4.2 RhoA partially activates mDia2
Previous studies using the in vitro actin polymerization assays have shown the auto-inhibition of formins is only partially relieved by RhoA, even in the presence of a vast molar excess of RhoA (Li and Higgs 2005; Maiti et al. 2012). Furthermore, the partial activation of mDia1 is independent of the nucleotide bound state in in vitro pyrene-actin polymerization assays; RhoA being equally potent in the GDP or GTP bound state (Li and Higgs 2005). I assessed the ability of RhoA to activate mDia2 using the pyrene-actin polymerization assay. RhoA(Q63L)-GTP (a constitutively activated form of RhoA, Longenecker et al. 2003) and
RhoA-GDP were added to the NT1-533-CT complex and the extent of the actin polymerization was analyzed. A 10 fold molar excess of RhoA in either the GTP or GDP bound state only weakly activated the actin polymerization activity of CT in the presence of NT1-533 (Fig 4.13A-C).
These data are consistent with our biochemical binding assays that demonstrate RhoA only partially disrupts the interaction between NT1-533-CT of mDia2. Furthermore, these data suggest that mDia1 and mDia2 are regulated in a similar manner by RhoA. More importantly these data suggest that other factors must be involved in the complete activation of formins, either in addition to or instead of RhoA.
74 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.12
NT1-533 inhibits the actin polymerization activity of CT. (A) In vitro actin polymerization reaction in the presence of increasing concentration of CT. (B) In vitro actin polymerization reaction in the presence of CT and increasing concentrations of NT1-533. (NT=NT1-533). (C)
Quantitation of the actin polymerization rate at t(1/2) (discussed in 6.5) in each group. The rate of actin by itself was set to 1.0 as control. Each individual group was repeated three times. For this set of experiments, two different batches of pyrene-actin/actin were used and proteins including CT, NT1-533 were freshly purified each time. Error bars indicate ± SEM.
75 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.13
RhoA weakly activates mDia2. (A)(B) In vitro actin polymerization reaction in the presence of the CT-NT1-533 complex and increasing concentration of RhoA (RhoA=RhoA(Q63L)-GTP;
NT=NT1-533). (C) Quantitation of the actin polymerization rate at t(1/2) in each group. The rate of actin polymerizaition by itself was set to 1.0 as control. Each individual group was repeated three times. Error bars indicate ± SEM.
76 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
4.2.4.3 Anillin has no effect on mDia2 actin polymerization activity
To determine if anillin has a role in activating mDia2 I performed a mDia2 actin polymerization assays in the presence of increasing concentrations of anillin. Anillin was added with the NT1-533-CT complex to analyze the extent of the actin polymerization. Even at
10-fold molar excess anillin did not disrupt the auto-inhibition of NT1-533-CT (Fig 4.14A,B).
These data support my biochemical binding results that anillin cannot bind to the NT1-533 in the presence of CT.
4.2.4.4 Anillin enhances RhoA dependent activation of mDia2 actin polymerization activity
Biochemical analysis of the role of RhoA and anillin in modulating the interaction between the NT and CT of mDia2 suggested that anillin functioned to enhance RhoA mediated disruption of the auto-inhibitory interaction between the NT and CT of mDia2 (Fig 4.8). These data suggest a model whereby anillin enhances the activation effect of RhoA on mDia2. To test this model actin polymerization assays were carried out in the presence of anillin,
RhoA and the NT1-533-CT complex. Anillin and RhoA were added to the NT1-533-CT complex to analyze the extent of the actin polymerization. In the presence of constant anillin, increasing concentrations of RhoA stimulated a small dose dependent rise in actin polymerization activity of mDia2 (Fig 4.15A,D). In contrast, increasing the concentration of anillin in the presence of a constant concentration of RhoA had no dose dependent activation effect (Fig 4.15B,D). However, upon increasing the concentration of anillin and RhoA, mDia2 activation increased significantly (Fig 4.15C,D). Combined, the biochemical data and the actin polymerization data suggest anillin acts to enhance RhoA dependent activation of mDia2.
77 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.14
Anillin does not activate mDia2 actin polymerization activity. (A) In vitro actin polymerization reaction in the presence of the CT-NT1-533 complex and increasing concentrations of anillin. (Ani=anillin; NT=NT1-533). (B) Quantitation of the actin polymerization rate at t(1/2) in each group. The rate of actin by itself was set to 1.0 as control. Each individual group was repeated three times. Error bars indicate ± SEM.
78 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.15
Anillin enhances the activation effect of RhoA on mDia2. (A) In vitro actin polymerization reaction in the presence of the CT-NT1-533 complex, constant concentration of anillin and increasing concentrations of RhoA (RhoA=RhoA(Q63L)-GTP; Ani=anillin; NT=NT1-533). (B)
In vitro actin polymerization reaction in the presence of the CT-NT1-533 complex, constant concentration of RhoA and increasing concentrations of anillin. (C) In vitro actin polymerization reaction in the presence of the CT-NT1-533 complex and increasing concentrations of anillin and RhoA. (D) Quantitation of the actin polymerization rate at t(1/2) in each group. The rate of actin by itself was set to 1.0 as control. Each individual group was repeated three times. Error bars indicate ± SEM.
79 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
4.2.5 The anillin-mDia2 interaction is required for modulating mDia2 activity
To confirm that a direct interaction between anillin and mDia2 is required for anillin mediated enhancement of mDia2 activity I sought to specifically disrupt the anillin-mDia2 interaction. I had mapped the mDia2 binding region on N-terminal of anillin between amino acids 22-57 (Fig 4.4). Sequence alignments of anillin from different vertebrate species identified two conserved regions in this section of anillin; N1 that extended from amino acid 1 to 32 and included a CD2AP binding site (Monze et al. 2005) and N2 from amino acids 38 to 44 (Fig 4.3). I demonstrated that CD2AP binding does not compete with mDia2 binding (Fig 4.5). These data suggested that mDia2 bound to a distinct site on anillin and the most likely site would be the N2 region between amino acids 38-44 (Fig 4.3). To test this model I generated a series of mutations between amino acid residues 38 and 44 converting single or multiple residues to alanine (Fig 4.16A). Anillin fragments containing these mutations were fused to MBP and analyzed for binding to NT89-533 of mDia2 (Fig 4.16B, right panel). Mutating KR (38-39)-AA had no significant effect on anillin binding to mDia2 (Fig 4.16B, Fig 4.17). Moderate effects on binding were observed in the mutants R41A (44±6.0% reduction), Q42A (14±5.0% reduction) and PL (43-44)-AA (61±6.0%) reduction (Fig 4.16B,Fig 4.17). However, the RQPL (41-44)-AAAA mutant reduced binding to mDia2 by 97±1.0% (Fig 4.16B, Fig 4.17). To determine if the RQPL (41-44)-AAAA mutant is specific for abrogating mDia2 binding, I analyzed its binding to CD2AP and importin β2. Anillin RQPL (41-44)-AAAA mutant bound equally well to CD2AP and importin β2 (Fig 4.16B, left panel) (the binding has not been compared to wild-type anillin binding to CD2AP and importin β2), suggesting the RQPL (41-44)-AAAA mutant uniquely disrupts the anillin- mDia2 interaction.
To directly test if the anillin-mDia2 interaction was required to enhance RhoA mediated mDia2 actin polymerization activity, the anillin mutant RQPL (41-44)-AAAA was utilized in the mDia2 actin polymerization assay. The anillin mutant and RhoA were added to the
NT1-533-CT complex to analyze the extent of the actin polymerization. In contrast to the wild-type anillin, the RQPL (41-44)-AAAA mutant did not enhance RhoA mediated activation of mDia2 (Fig 4.18A, B). These data suggest a direct interaction between anillin and mDia2 is required for modulating mDia2 activity.
80 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.16
In vitro binding assays reveal anillin residues that contribute to mDia2 binding. (A) Cross-species sequence alignments showing proposed mDia2 binding sites. Anillin mutants containing different amino acids mutations are marked with arrows below. Numbers correlate to those used in (B). (B) Western-blotting analysis showing pull down anillin mutants by
NT89-533. The bands of anillin are indicated by arrows.
81 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.17
Quantitation of blot intensities of the pull down groups in Fig 4.16. Blot intensity of wild-type pull down group was set to 1.0 and blots intensities of mutant groups were compared to the wild-type pull down group. Three parallel experiments were repeated. The intensities of wild-type pull down blots were set to 1.0. Error bars indicate ± SEM.
82 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.18
The anillin-mDia2 interaction is required for modulating mDia2 activity. (A) In vitro actin polymerization reaction in the presence of the CT-NT1-533 complex and increasing concentrations of the anillin mutant and RhoA (RhoA=RhoA(Q63L)-GTP; Anlnmut=anillin mutant RQPL-AAAA; NT=NT1-533). (B) Quantitation of the actin polymerization rate at t(1/2) in each group. The rate of actin by itself was set to 1.0 as control. Each individual group was repeated three times. Error bars indicate ± SEM.
83 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
4.3 Discussion and future directions
In this study, I identified the anillin-mDia2 interaction and mapped the mDia2 binding site on anillin. My results demonstrate that, anillin by itself has no effect, but enhances RhoA inhibition of the mDia2 DID-DAD autoinhibitory interaction. Using in vitro actin polymerization assays, I demonstrate that anillin enhances RhoA dependent activation of mDia2 actin polymerization activity. My data addresses a puzzle within the actin field of how formins are fully activated at the correct time and place within the cell. Most previous models have been built upon G-protein dependent activation of formins (Wallar and Alberts 2003; Seth et al. 2006). However, both in vivo and in vitro data suggested these models were an oversimplification of the data (Palazzo et al. 2004; Li and Higgs 2005; Higashi et al. 2010; Maiti et al. 2012). My results provide a model to resolve these issues (Fig 4.19A). In this model, G-proteins like Rho first interact with the GBD domain of formins and relieve the DAD-DID autoinhibitory binding. Subsequently an enhancer binds the DID domain and completely displaces DAD from DID, leading to full activation of formins (Fig 4.19A). In the case of the anillin-mDia2 interaction in cytokinesis, this molecular model would ensure that mDia2 is only recruited and fully activated at the sites of the plasma membrane where anillin and RhoA are enriched, the site of the cleavage furrow ingression. Moreover, the model describes a novel mechanism of how cytoskeletal actin nucleation is regulated that can be applied throughout the formin family. This model suggests that small G-proteins, in conjunction with DID domain binding proteins, combined to spatially regulate the site of actin nucleation within a cell.
There is mounting evidence identifying different ligands that bind to formin regions that include the DID domain (Fig 4.19B) (Chesarone et al. 2010). For example IQGAP1 binds to the DID domain of mDia1 and competes with DAD domain binding (Brandt et al. 2007). This mechanism is reminiscent of the way anillin interacts with mDia2. In mammalian cells IQGAP is required for mDia1 localization to the phagocytic cup. Its role in modulating mDia1 activity is unclear. In the next step I will test if other DID-binding ligands (such as listed in Fig 4.19B) have similar regulatory roles as anillin. Such a mechanism would allow the tight spatial and temporal activation and formins to govern precise control of the
84 Chapter 4 Anillin enhances RhoA dependent activation of mDia2 organization of the actin cytoskeleton.
Using in vitro binding assays I identified the RQPL (41-44) residues of anillin as the mDia2 binding site. These residues are conserved among vertebrate anillin sequences but are not presented in Drosophila anillin (Fig 4.16A). This may explain the results from previous studies in Drosophila S2 cells where anillin does not have a significant role early in cytokinesis, rather it has later roles in furrow stabilization and ICB formation (Echard et al. 2004; Hickson and O’Farrell 2008). One possibility is that Drosophila S2 cells utilize another mechanism, other than the anillin-mDia2 interaction, to recruit formins to the membrane furrow at the early stage of cytokinesis.
Interestingly, the RQPL (41-44) residues are also involved in the ubiqutination of anillin. Anillin is a substrate of the anaphase-promoting complex/cyclosome (APC/C) (Zhao and Fang 2005). The level of expression of anillin is cell-cycle dependent, peaking in mitosis and dropping in G1 (Field and Alberts 1995). Anillin is degraded during mitotic exit by the APC/CCdh1 complex through a destruction-box (D-box) RQPL (41-44) (Zhao and Fang 2005). Interestingly, I identified these residues as being required for mDia2 binding (Chapter 4.2.5). It raises an interesting question if ubiqutination being involved in regulating the anillin-mDia2 interaction. In late anaphase and telophase, mDia2 was reported to be concentrated at the ICB yet its roles at the ICB were undefined (Wanatabe et al. 2008). Meanwhile the Wilde lab showed that anillin concentrated at the furrow when the ICB matured and was gradually depleted from the ICB after relocating to the sites of microtubule constriction (Renshaw et al. 2014). To identify the role of anillin-mDia2 interaction in cytokinesis and determine if the ubiquination is involved in regulating it, I will establish stable cell lines that express GFP-anillin (RQPL-AAAA) mutant and use anillin RNAi-rescue experiment to examine the mutant phenotypes. The mutant phenotype will reveal when and where the anillin-mDia2 interaction is required for cytokinesis.
My data demonstrate that anillin is not able to interact with NT1-533 and CT complex unless RhoA is present. In contrast, RhoA by itself can interact, although weakly, with the
NT1-533-CT complex. These results highlight how these proteins interact with each other in time and space; that RhoA is required to bind to mDia2 first to facilitate the binding of anillin, leading to a full activation of mDia2. Previous crystallography studies have suggested three
85 Chapter 4 Anillin enhances RhoA dependent activation of mDia2 ways that Rho binding to GBD occludes the DAD from the DID to release the autoinhibitory DAD-DID binding of mDia (Lammers et al. 2005; Nezami et al. 2006). RhoA binding to GBD can change the GBD-DID domains to a closed conformation where GBD occludes the DAD peptides from the DAD-binding pocket of DID (Lammers et al. 2005). In addition, the residue Arg68 of RhoA sterically interferes with DAD peptide. Residues Glu64 and Asp65 of RhoA that project toward the acidic residues on the surface of DAD peptide also electrostatically repulse the DAD (Lammers et al. 2005; Nezami et al. 2006). Based on these results I propose a model that before RhoA binds to mDia2, the anillin binding surface on DID is occupied by the DAD. Upon RhoA binding, the RBD (and RhoA by itself) occludes DAD from the DID to make space for the anillin-DID interaction. Subsequently, anillin binding to the DID accelerates the displacement of DAD from DID. Together anillin and RhoA form a complex with mDia2 to stabilize the activated state of mDia2.
86 Chapter 4 Anillin enhances RhoA dependent activation of mDia2
Fig 4.19
A new model of cellular formin activation and a list of DID-binding ligands. (A) Cartoon of an enhancer required, G-protein dependent formin activation model. In the cases when G-proteins are not sufficient to activate formins, enhancers provide localized activities for formin activation and participate in modulating the actin cytoskeleton. (B) A list of DID-binding ligands. Asterisks indicate two mammalian DID-binding ligands.
87 Summary
5. Summary
In this study I’ve identified multiple roles of N-terminal region of anillin. First I identified a NLS that targeted anillin to the nucleus in interphase by binding to importin β2. The NLS was mapped out between N-terminal 1-91 amino acids of anillin and bound to importin β2 via a typical basic patch- (PY) motif. Although the anillin-importin β2 interaction had no role in cytokinesis, the nuclear targeting of anillin in interphase was required to maintain normal cell shape.
Using confocal microscopy I was able to identify anillin localized to the basal body of cilia in differentiated cells. The result of anillin RNAi suggested anillin could be involved in cilia morphogenesis.
I mapped the binding site of mDia2 on anillin and discovered that the anillin-mDia2 interaction enhances RhoA dependent activation of mDia2 actin assembly activity. I determined the binding site of mDia2 was distinct from those of CD2AP and importin β2. I demonstrated later that the anillin-mDia2 interaction was critical for modulating mDia2 activity. Furthermore I established that RhoA binding to mDia2 facilitated subsequent anillin binding. Based on these findings I proposed a new model for formin activation whereby a DID binding factor enhances small G-protein activation of formin. Such a model reasonably addresses the issues of how formins are activated at the right time and location within cells. In the case of anillin, this suggest it is not simply an actin binding protein to cross-link actin filaments (Kinoshita et al. 2002), but also involved in regulation of the actin filament production via modulating mDia2 activity.
88 Materials and methods 5.2 cDNA constructs generation
6. Materials and methods
6.1 Materials
6.1.1 Bacterial strains
The E.coli strain, TOP 10, was used for molecular cloning and the propagation of plasmids. The genotype of the strain is: F-mcrAΔ (mrr-hsdRMS-mcrBC) ϕ80lacX74 recA1 araD139 Δ(ara-leu)7697 galU galK rpsL (StrR) endA1 nupG.
The BL21 E.coli strain was used for the expression of GST- or 6xHis- tagged proteins. The genotype of the strain is: fhuA2 [lon] ompT gal [dcm] ΔhsdS.
The ER2523 E.coli strain was used for the expression of MBP- tagged proteins. The genotype of the strain is: fhuA2 [lon] ompT gal sulA11 R(mcr-73::miniTn10--TetS)2 [dcm] R(zgb-210::Tn10--TetS) endA1 Δ(mcrC-mrr)114::IS10.
6.1.2 Kits used See Table 6.1 in Supplementary documents.
6.1.3 Nucleic acids
See Table 6.2, Table 6.3 in Supplementary documents
89 Materials and methods 5.2 cDNA constructs generation
6.2 cDNA constructs generation
6.2.1 DNA amplification and isolation
For amplification of full-length cDNA or fragments, template DNA was selectively amplified by polymerase chain reaction (PCR) using the i-Max IITM DNA polymerase (Froggalab). The 5’ and 3’ oligonucleotide primers were manually designed for each cDNA sequence and were ordered from IDT (Coralville, Iowa) (Table 5.5-5.6). PCR were carried out in a PTC-200 thermal cycler (MJ Research). The following components are added into reactions to a final concentration of: 10-25pg/µl DNA template, 0.5µM primers (5’ and 3’), 0.2mM dNTP mixture, 1x reaction buffer (Froggalab), 0.25% (v/v) i-Max II DNA polymerase. Briefly, the template DNA was denatured at 95°C for 30s, followed by primer annealing at 55-60°C for 30s and primer extension at 72°C. In total, 30 cycles of PCR were repeated before the reaction was stopped. Amplified DNA products were further isolated and purified by the Gel/PCR DNA Fragment Extraction Kits (Geneaid) following the manufacturer’s instructions.
For the generation of mutations, two primers with overlapping complimentary ends that contain the mutations were mixed separately with their paired initial (5’) and (3’) primers (IDT, Coralville, Iowa) (Table 6.3). PCR was first carried out to generate two DNA products with overlapping ends that encompass the mutations. Then the two DNA products were equally mixed to a final concentration of 25-50pg/µl and amplified by PCR for 10 cycles without primers. Afterwards, the initial (5’) and end (3’) primers were added into the reaction each to a final concentration of 0.5µM and the reaction was continued for another 20 cycles. Amplified DNA products were further isolated and purified using the same method described above.
After isolation and purification, DNA fragments were analyzed on 1% agarose gel dissolved in 1xTAE buffer (40mM Tris-acetate pH=8.0, 1mM EDTA) with 0.5µg/ml ethidium bromide for DNA visualization. DNA samples were mixed with 1xDNA loading buffer (1.7mM Tris-HCl pH=7.6, 0.005% bromophenol blue, 0.005% xylene cyanol FF, 10% glycerol,
90 Materials and methods 5.2 cDNA constructs generation
10mM EDTA) (Fermentas) and loaded onto agarose gels. DNA was separated by gel electrophoresis. The concentration of DNA was determined spectrophotometrically at A260 using a BioMate-3 spectrophotometer (Thermo Scientific).
6.2.2 Cloning in PCR8/GW/TOPO system
Amplified cDNA products were isolated and purified as described above, then A-tailed using Taq DNA polymerase (Fermentas) in the presence of 2.5mM dATP (Fermentas) at 72°C for 15-20min. The A-tailed DNA fragment was subcloned into the PCR8/GW/TOPO vector (Invitrogen) following the manufacturer’s instructions. The correct orientation of the cDNA in the plasmid was determined by restriction mapping.
To generate plasmids that expressed MBP (N-) tagged recombinant proteins, the cDNA fragments was sub-cloned from the PCR8/GW/TOPO vectors by recombination into the pKM596 (Addgene) vector using the LR Clonase™ II Enzyme Mix (Invitrogen) following the manufacturer’s instructions. To generate plasmids that expressed GST (N-) or the 6xHis (N-) tagged recombinant proteins, the cDNA fragments in PCR8/GW/TOPO were recombined into the pDEST 15 or pDEST 17 vectors (Invitrogen), respectively, using the LR Clonase™ II Enzyme Mix following the manufacturer’s instructions. To generate plasmids that were to be used for transient transfection in HeLa cells to express GFP (N-)-anillin fusion proteins, the anillin cDNA fragments in PCR8/GW/TOPO were recombined into the pcDNA6.2/N-EmGFP- DEST vector using the LR Clonase™ II Enzyme Mix following the manufacturer’s instructions. Plasmid DNA was sequenced by ACGT (Toronto, ON) to confirm that the cDNA sequence was in-frame and contained no spurious mutations.
6.2.3 Ligation independent cloning (LIC) system
To generate eGFP-anillin constructs that stably express in cell lines, cDNA products were amplified using primers that contained sequences for LIC cloning (Table 6.2). cDNA products were isolated and purified as described above. Purified cDNA products were treated with T4 DNA polymerase (Invitrogen) to generate single stranded overhangs on 5’ and 3’ ends. The following components are added into reaction in a final concentration of: 10-50ng/µl of DNA, 5%(v/v) T4 DNA polymerase, 1x T4 polymerase reaction buffer
91 Materials and methods 5.2 cDNA constructs generation
(Invitrogen), 2.5mM dTTP, 5mM DTT. The reaction was incubated at room temperature for 30min and stopped at 75°C for 20min with the presence of 10mM EDTA. The pcDNA5/FRT/TO/eGFP-N vector was previously modified to contain the LIC sequence (Liu et al. 2012; Renshaw et al. 2014). The vector (1-2µg) was linearized by Fse I (New England Biolabs) at 37°C for 60min and Fse I was heat inactivated at 65°C for 20min. The vector was then treated with T4 DNA polymerase as described above but in the presence of 2.5mM dATP instead of dTTP to generate single stranded overhangs at the 5’ and 3’ ends. To anneal the cDNA to the vector, cDNA and vector were mixed and incubated for 10min at room temperature with presence of 40mM EDTA. The reaction was heated to 75°C and cooled slowly to room temperature and then transformed into TOP10 cells. Plasmid DNA was sequenced by ACGT to confirm that the cDNA sequence was in-frame and contained no spurious mutations.
92 Materials and methods 5.3 Protein expression and purification
6.3 Protein expression and purification
6.3.1 Bacterial transformation
For bacterial transformation, 50µl-100µl of thawed competent cells was incubated with plasmids on ice for 30min. The mixture was heat-shocked at 42°C for 90s then quickly ice-cooled for 120s before S.O.C media (Invitrogen) was added. The cell suspension was then incubated at 37°C for 60min (120min for spectinomycin-resistance selection). Cells were then plated on LB plate containing selective antibiotics in a final concentration of 100mg/ml ampicillin or 100mg/ml spectinomycin).
6.3.2 Protein expression
BL21 cells were grown in LB media at 37°C to an optical density of 0.6 at 600nm. Recombinant protein expression was then induced by the addition of 1mM IPTG (iso-propyl β-D-1-thiogalactopyranoside) and the cells were incubated at 16°C overnight, shaking at 180-225 rpm. Cells were then harvested by centrifugation at 7000g for 20min. The cell pellet was stored at -80°C while awaiting protein purification.
6.3.3 Protein purification
6.3.3.1 MBP (N-) fusion proteins purification
For the purification of MBP (N-) fusion proteins, E.coli (ER2523) cell pellets were resuspended in MBP/GST-lysis buffer containing 25mM HEPES pH 7.5, 500mM NaCl, 100mM KCl, 0.5mM 2-β-mercaptoethanol, 1mM PMSF and completely lysed by sonication. Cell lysates were then clarified by centrifugation at 10000g for 30min at 4°C to isolate supernatant from the cell fragment and precipitant. The supernatant was applied to amylose resin (New England Biolabs) following the manufacturer’s instruction. Briefly, the amylose resin was equilibrated in MBP/GST-CB containing 25mM HEPES pH 7.5, 500mM NaCl, 100mM KCl, 0.5mM 2-β-mercaptoethanol, 1mM PMSF, 0.1% (v/v) Triton X-100. Then the
93 Materials and methods 5.3 Protein expression and purification supernatant was run over through amylose resin thoroughly and the amylose resin was washed with 10 column volumes of MBP/GST-CB 3 times. The MBP-proteins were finally eluted by 10mM maltose dissolved in MBP/GST-CB.
6.3.3.2 GST (N-) fusion proteins purification
For the purification of GST (N-) fusion proteins, E.coli cell pellets were lysed and the supernatant was collected as described above. The supernatant was then applied to glutathione agarose column (Fermentas) following the manufacturer’s instructions. Briefly, the glutathione agarose was equilibrated in MBP/GST-CB and the supernatant was run over. The column was then washed with 10 column volumes of MBP/GST-CB 3 times. The GST-proteins were finally eluted by 10mM glutathione in MBP/GST-CB.
6.3.3.3 6xHis (N-) fusion proteins purification
For the purification of 6xHis (N-) fusion proteins, E.coli cell pellets were resuspended in 6xHis-lysis buffer containing 25mM HEPES pH 7.5, 500mM NaCl, 5% (v/v) Glycerol, 5mM imidazole, 0.5mM 2-β-mercaptoethanol, 1mM PMSF and completely lysed by sonication. Cell lysates were then clarified by centrifugation at 10000g for 30min at 4°C to separate soluble material from insoluble material. The supernatant was applied to Ni-sepharose beads (Amersham Biosciences) following the manufacturer’s instructions. Briefly, the Ni-sepharose beads were equilibrated in 6xHis-CB containing 25mM HEPES pH 7.5, 500mM NaCl, 5% (v/v) Glycerol, 20mM imidazole, 0.5mM 2-β-mercaptoethanol, 1mM PMSF. Then the supernatant was applied to the Ni-sepharose column. The beads were washed with 10 column volumes of 6xHis-CB 3 times. The 6xHis-proteins were finally eluted by 500mM imidazole dissolved in 6xHis-CB.
6.3.4 Dialysis and storage
After elution, the peak protein fractions were pooled and dialyzed against 10mM HEPES pH
7.6, 100mM KCl, 2mM MgCl2, 0.1mM CaCl2, 50mM sucrose, 5mM EGTA overnight at 4°C then concentrated in a Milipore Ultrafree Spin Device (10K MWCO) (MILIPORE Ireland) by centrifugation at 6000rpm for 10min in 4°C. Proteins for in vitro binding assays were
94 Materials and methods 5.3 Protein expression and purification aliquoted and flash frozen in liquid nitrogen prior to storage at -80°C. The concentration of purified proteins was measured using a BioRad protein assays and by running SDS-PAGE gels of serially diluted purified proteins and BSA. Gels were stained with Coomassie and quantified by Quantity One Software to compare the concentration of purified proteins to the concentration of standard protein BSA.
6.3.5 Nucleotide loading
To generate GTP or GDP loaded GST-RhoA for the in vitro binding and actin polymerization assays, the peak GST-RhoA protein fractions were pooled after elution from the glutathione agarose column. Protein concentrations were determined using the BioRad protein assays as described above. Proteins were then added to 25mM EDTA, 1mM DTT and GTP or GDP added to a 100x molar excess of the proteins. The reactions were incubated on ice for 40min before adding MgCl2 to a final concentration of 50mM. Proteins were then further dialyzed against 100mM Hepes pH 7.7, 100mM KCl, 2mM MgCl2, 50mM sucrose, 5mM EGTA overnight 4°C, concentrated, aliquoted then flash frozen in N2 (l) and stored at -80°C as described above.
95 Materials and methods 6.4 In-vitro binding assays
6.4 In vitro binding assays
6.4.1 Anillin and importins binding assays
To test the ability of wild-type or mutant fragments of anillin to bind to importin β1 or β2, 0.05nmol of purified MBP-anillin fragments were immobilized onto 20µl of amylose resin beads in 100µl of incubation buffer (50mM Hepes pH 7.5, 50mM NaCl, 1mM EDTA, 5mM
MgCl2, 0.3% (v/v) Triton X-100, 1mM 2-β-Mercaptoethanol), for 1h at 4°C. The resin was washed with incubation buffer 3 times then blocked with 3% (w/v) BSA for 20min. The resin was washed three times with incubation buffer then mixed with 0.02nmol of purified 6xHis-importin β1/β2 and incubated for 4h at 4°C. For RanGTP competition assay, 0.05nmol of purified MBP-anillin fragments were immobilized onto amylose resin beads and the resin was washed then blocked as described above. The resin was mixed with 0.02nmol of purified 6xHis-importin β1/β2 and incubated for 4h at 4°C. The resin was washed 3 times with incubation buffer and 0.02nmol RanGTP was added to incubate for 2h at 4°C. The resin was then washed three times with incubation buffer, isolated by centrifugation and boiled in SDS sample buffer then analyzed by Western-blotting using an anti-6xHis polyclonal antibody to detect co-purifying importins (MP Biomedicals).
To map where importin β2 bound on the N-terminus of anillin, 0.05nmol of purified 6xHis-importin β2 was immobilized onto 10µl Ni-Sapharose beads (Amersham Biosciences) in 100µl of incubation buffer (25mM Hepes pH 7.5, 120mM NaCl, 1mM EGTA, 0.3% (v/v) Triton X-100, 1mM 2-β-Mercaptoethanol), for 1h at 4°C. The beads were washed 3 times with the incubation buffer and blocked with 3% (w/v) BSA for 20min. The beads were washed three times with incubation buffer and mixed with 0.02nmol of purified MBP-anillin fragments and incubated for 4h at 4°C. The beads were then washed three times with incubation buffer, re-isolated by centrifugation and boiled in SDS sample buffer then analyzed by Western-blotting using an anti-MBP monoclonal antibody to detect co-purifying anillin fragments (New England Biolabs).
96 Materials and methods 6.4 In-vitro binding assays
6.4.2 Anillin and mDia2 binding assays
To map out the anillin-mDia2 binding, 0.05nmol of purified GST-NT89-533 or CT was immobilized onto 20µL glutathione agarose in 100µl of incubation buffer including 50mM Hepes pH 7.5, 50mM NaCl, 1mM EDTA, 5mM MgCl2, 0.3%(v/v) Triton X-100, 1mM β-Mercaptoethanol and incubated for 1h at 4°C. The beads were washed and blocked with 3% (w/v) BSA for 20min. The resin was then washed for three times with incubation buffer and mixed with 0.02nmol of purified different MBP-anillin fragments and incubated for 2h at 4°C. The beads were then washed three times with incubation buffer, re-isolated by centrifugation and boiled in SDS sample buffer then analyzed by Western-blotting using an anti-MBP monoclonal antibody to detect co-purifying anillin fragments (New Englands Biolabs).
6.4.3 Anillin, RhoA and mDia2 competition assays
To test the competition effects of anillin and Rho on the interaction between NT1-533 and CT,
0.05nmol purified MBP-NT1-533 was immobilized onto 20µl amylose resin in 100µl of incubation buffer as described above and incubated for 1h at 4°C. The resin was washed 3 times with incubation buffer then blocked with 3% (w/v) BSA for 20min. The resin was washed 3 times with incubation buffer, then mixed with 0.05nmol purified GST-CT and incubated for 2h at 4°C. The resin was washed 3 times with incubation buffer and 6xHis-anillin and GST-RhoA were added to incubate for 2h at 4°C. The resin was washed 3 times with incubation buffer, re-isolated by centrifugation and boiled in SDS sample buffer then analyzed by Western-blotting using a homemade anti-GST polyclonal antibody to detect co-purifying GST-CT. To determine if anillin or Rho bind to the CT-NT1-533 complex first, the same protocol was used, except analysis was carried out with an anti-His polyclonal antibody (MP Biomedicals) to detect co-purifying anillin and the homemade anti-GST polyclonal antibody to detect co-purifying Rho.
6.4.4 Importin β2, CD2AP and mDia2 competition assays
To determine if importin β2 and mDia2 shared a common binding site on anillin, 0.05nmol of purified MBP-ANLN (1-151) was immobilized onto 20µl of amylose resin beads in 100µl of incubation buffer (50mM Hepes pH 7.5, 50mM NaCl, 1mM EDTA, 5mM MgCl2, 0.3% (v/v)
97 Materials and methods 6.4 In-vitro binding assays
Triton X-100, 1mM 2-β-Mercaptoethanol), for 1h at 4°C. The resin was washed 3 times with incubation buffer and blocked with 3% (w/v) BSA for 20min. The resin was washed three times with incubation buffer then mixed with 0.02nmol of purified GST-NT89-533 and incubated for 2h at 4°C. The resin was washed three times with incubation buffer and added with different concentrations of 6xHis-importin β2 to incubate for 4h at 4°C. The resin was washed three times with incubation buffer, re-isolated by centrifugation and boiled in SDS sample buffer the analyzed by Western-blotting using a homemade anti-GST polyclonal antibody to detect co-purifying mDia2.
To determine if CD2AP and mDia2 shared a common binding site on anillin, 0.05nmol of purified MBP-ANLN (1-151) was immobilized onto 20µl amylose resin beads in 100µl of incubation buffer (50mM Hepes pH 7.5, 50mM NaCl, 1mM EDTA, 5mM MgCl2, 0.3% (v/v) Triton X-100, 1mM 2-β-Mercaptoethanol), for 1h at 4°C. The resin was washed with incubation buffer and blocked with 3% (w/v) BSA for 20min. The resin was washed 3 times with incubation buffer then mixed with 0.02nmol of purified GST-NT89-533 and incubated for 2h at 4°C. The resin was washed 3 times with incubation buffer and added with different concentrations of GST-CD2AP to incubate for 2h at 4°C. The resin was washed 3 times with incubation buffer, re-isolated by centrifugation and boiled in SDS sample buffer then analyzed by Western-blotting using a homemade anti-GST polyclonal antibody to detect co-purifying mDia2.
98 Materials and methods 6.5 Quantitation of blots intensities
6.5 Quantitation of blots intensities
Western blots were visualized using BioRad MP Imager (Bio-Rad Canada). Only images that were not overexposed were selected for quantitative analysis. Specific bands in specific lanes were selected using ‘Set up lanes and bands’ functions in the ImageLab software (Bio-Rad Inc.). Using the ‘Relative Comparison’ function in ‘Quantitation Tools’ of the software, the control bands of the pull-down groups can be selected and the value of blot intensity of the control band was set to 1.0. Using the ‘Analysis Diagram’ function of the software, the relative values of blot intensities of all other bands in pull-down groups can be automatically quantified by the software (by comparing their values to the control band).
Meanwhile, using the ‘Relative Comparison’ function of the software, the control bands of the loading control groups can be selected and the value of blot intensity of the control band was set to 1.0. Using the ‘Analysis Diagram’ function of the software, the relative values of blot intensities of all other bands in loading control groups can be automatically quantified by the software.
To normalize the blot intensities of the pull down groups, the intensity value of each pull down group was divided by the intensity value of each corresponding loading control group. The resulted value was the normalized blot intensity for each pull down group (relative to the control band of the pull down group).
99 Materials and methods 6.6 In vitro actin polymerization assay
6.6 In vitro actin polymerization assays
To assess the actin polymerization activity of mDia2, lyophilized pyrene-labeled and unlabeled actin (Cytoskeleton) were first resuspended in ddH2O and equilibrated in G-buffer
(monomer actin buffer) including 2mM Tris pH 8.0, 0.2mM CaCl2, 0.2mM ATP, 0.5mM β-Mercaptoethanol on ice for 2h. The pyrene-labeled actin was mixed with un-labeled actin in a 15%: 85% ratio. Proteins were added to actin and incubated for 5-10min at room temperature. To initiate the reaction, the mixture was treated with polymerization buffer including 25mM Tris pH 7.0, 50mM KCl, 2mM MgCl2, 0.1mM ATP. The increase in fluorescence intensity was monitored in a PTI fluorimeter with excitation at 365nm and emission at 386nm as previously described (Arora and McCulloch 1996).
The methods of measuring the actin polymerization rate of each curve were described before (Doolittle et al. 2013). To quantify all of the curves more efficiently while maintaining the statistical accuracy, I modified the methods as following steps (Fig 6.1):
(1). Pick up 10 data points at the minimum/maximum intensities to calculate Imin (the average minimum intensity) and Imax (the average maximum intensity). The number of the sampled points can be adjusted according to the flatness of the specific curve.
(2). Pick up data points with intensities between 0.48×(Imax-Imin)+Imin and
0.52×(Imax-Imin)+Imin and visually check to confirm these points fall in the middle of the dataset of I values.
(3). Fit a linear line to these points and visually check if the line fits to most of the sampled points. Record the slope and intercept of the line as m(1/2) and b(1/2), respectively.
(4). t(1/2) (time point when half amount of total actin monomers polymerize to filaments) of the curve can be calculated as:
t(1/2) =(0.5×(Imax - Imin)+ Imin + b(1/2))/ m(1/2)
100 Materials and methods 6.6 In vitro actin polymerization assay
e. APt(1/2) (actin polymerization rate at t(1/2)) of the curve can be calculated as:
APt(1/2) = 1.88×m(1/2)/(Imax - Imin)
f. To confirm the accuracy of the calculation, the [Actin](t1/2) (the remaining free actin at t(1/2))can be determined by:
[Actin](t1/2) = [Actin](t0)- (APt(1/2)/ m(1/2)) ×((m(1/2) ×t(1/2)- b(1/2))- Imin)
[Actin](t0) means the initial actin monomer concentration in the reaction. In this study, the value of [Actin](t0) is 2.0(µM). Ideally, the [Actin](t1/2) value should be 1.0.
101 Materials and methods 6.6 In vitro actin polymerization assay
Fig 6.1
Graphical representation of the methods to analyze the in vitro pyrene actin polymerization data. Shown are the metrics of the data used to calculated the Imax and Imin， m(1/2) (slope of the linear fitted line in cyan), b(1/2) (intercept of the cyan line), t(1/2), actin polymerization rate at t(1/2) and the remaining free actin at t(1/2). (Figure adapted from Doolittle et al. 2013)
102 Materials and methods 6.7 Immunostaining and microscopy
6.7 Immunostaining and microscopy
6.7.1 Cell culture and transfection
HeLa cells were cultured in DMEM3 (Sigma) supplemented with 10% fetal bovine serum
(FBS) (Invitrogen) in a 5% CO2 atmosphere at constant temperature (37°C) and humidity. For transient transfection, plasmids expressing EmGFP-anillin fusion proteins were transfected into Hela cells grown on glass coverslips by using Lipofectamin® 2000 Reagent (Invitrogen). For one well of a 6-wells plate, add 2.5µg of DNA and 7.5µl of Lipofectamine 2000 into 250µl OptiMEM media (Sigma). Incubate the mixture in room temperature for 20min and then transfect the cells. 24 hours after transfection (48 hours for overexpression), cells were fixed in 3.7% paraformaldehyde (EMD) in PBS for 15 min at room temperature. Cells were then permeabilized in 0.1% Triton X-100 in PBS for 15min at room temperature. hTERT RPE-1 cells were culured in DMEM-F12 (Sigma) supplemented with 10% FBS
(Invitrogen) in a 5% CO2 atmosphere at constant temperature (37°C) and humidity. To induce differentiation, cells were serum starved in DMEM-F12 without FBS for 24-72 hours. To optimize the visualization of acetylated-tubulin, cells were fixed in -20°C methanol for 10 min. Cells can be fixed in 3.7% paraformaldehyde or 10% Trichloroacetic Acid (TCA) in PBS to visualize γ-tubulin.
6.7.2 Stable cell lines generation and characterization
The stable cell lines expressing eGFP-anillin and eGFP-anillin-KKR (68-70)-AAA from a Tet-inducible promoter were generated using the Flp-In system (Invitrogen) following the manufacturer’s instructions. Plasmids were first transfected into a HeLa Flp-In host cell line that contains an integrated FRT site and expresses the tetracycline repressor protein (TetR) by using Lipofectamin® 2000 Reagent (Invitrogen) (Renshaw et al. 2014). Transfected cells were selected using 200 µg/ml hygromycin B (BioShop) and 5 µg/ml blasticidin (Invitrogen). Monoclonal cell lines were selected from individual colonies that express eGFP-anillin or eGFP-anillin-KKR (68-70)-AAA and maintained in 200 µg/ml hygromycin B and 5 µg/ml
103 Materials and methods 6.7 Immunostaining and microscopy blasticidin. The expression of GFP-constructs can be induced by 25µg/ml of doxycycline for 24-72 hours prior to analysis.
6.7.3 siRNA treatment and rescue siRNA transfection to HeLa cells and subsequent rescue experiments were carried out as previously described (Renshaw et al. 2014, Liu et al. 2012). Briefly cells were transfected with 40nM double-strand anillin siRNA using Lipofectamin® 2000 Reagent (Invitrogen). 16-24 hours after siRNA treatment, cells were treated with doxycycline to induce the expression of eGFP-anillin and eGFP-anillin-KKR (68-70)-AAA. siRNAs were obtained from Integrated DNA Technologies. The 3’UTR siRNA duplexes for anillin (5’-agcuuacagacuuagcau-3’) and the negative control siRNA (5’-cguuaaucgcguauaauacgcgut -3’) were as previously described (Liu et al. 2012; Renshaw et al. 2014).
6.7.4 Immunofluorescence and microscopy
To visualize cells expressing different GFP (N-)-constructs, fixed cells then stained with 4', 6-diamidino-2-phenylindole (DAPI) to visualize DNA and an anti-α tubulin monoclonal antibody DM1A (Sigma) followed by a secondary goat anti-mouse antibody conjugated to Alexa 594 (Invitrogen) to visualize the cytosolic microtubule network. Alternatively cells were stained with rhodamine-labeled phalloidin (Invitrogen) to visualize the actin cytoskeleton network. Coverslips were mounted on glass slides using Mowiol (Polyvinyl alcohol 4-88, Fluka). Cells were visualized using a Nikon TE800 microscope with a 60x/1.4 NA oil-immersion objective lens and 1.515 immersion oil (Nikon) at room temperature. Images were acquired using METAMORPH software (Molecular Devices) driving an electron multiplying charge-coupled device (CCD) camera (ImagEM, Hammamatsu) as previously described (Renshaw et al. 2014). Z sections (0.2 µm apart) were acquired to produce a stack that was then imported into AUTOQUANT X2 (Media Cybernetics) for deconvolution (10 iterations). Maximum projections and cross sections were performed using METAMORPH. Images were overlaid in PHOTOSHOP (Adobe), involving adjustments in brightness and contrast of images.
104 Materials and methods 6.8 Statistical analysis
6.8 Statistical analysis
For all continuous data being quantified, individual experiments for each group were repeated at least 3 times and the means ± S.E were computed. When appropriate, the differences between groups were evaluated by Student’s unpair t test with statistically significance set at P<0.05.
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Supplementary documents
Table 6.1 Kits used in this work
Name Usage Provider Presto™ Mini Plasmid Kit DNA plasmid mini-prep Geneaid Biotech Gel/PCR DNA Fragments DNA isolation and gel Geneaid Biotech Extraction Kits extraction Geneaid Plasmid Midi Kit Plasmid midi-prep and Geneaid Biotech purification EZ-10 Spin Column DNA Gel DNA plasmid mini-prep, Bio Basic Extraction Kit isolation and gel extraction GenElute™ Gel Extraction Kit DNA plasmid mini-prep, Sigma isolation and gel extraction GenElute™ HP Plasmid Midi DNA plasmid midi-prep and Sigma Prep Kit purification PCR®/GW/TOPO® TA PCR products cloning Invitrogen cloning® Kit Lipofectamin® 2000 Reagent DNA and RNA transfection Invitrogen
116 Supplementary
Table 6.2 Oligonucleotides used to generate wild-type constructs
Name Sequence of oligos(5’!3’) Feature 5’_anln ATGGATCCGTTTACGGAGA 5’-initial forward primer of anillin
3’_anln TTAAGGCTTCCCAACAGGCTT 3’-end reverse primer of anillin
5’_anln_Lic GCTGGCGCTGGTGCGGGTGCC 5’-initial forward primer of anillin for GGAATGGATCCGTTTACGGA Lic entry cloning
3’_anln_Lic TCCGTGGCGGCCTCGTCGTCGG 3’-end reverse primer of anillin for Lic GATTAAGGCTTCCCAACAGGC entry cloning fwd_anln_22 AAAATGGCTGAGAGGCCCA 5’- forward primer for generating ANLN (22-151aa) fwd_ anln _57 GAGAAATCTTGTACAAAACC 5’- forward primer for generating ANLN (57-91aa); ANLN (57-151aa) fwd_ anln _91 GCAAAATCTTGTTCTCCA 5’- forward primer for generating ANLN (91-151aa); ANLN (91-450aa) rev_ anln _57 TTCACCACCAGAGAGGGG 3’- reverse primer for generating ANLN (1-57aa) rev_ anln _91 TGTCGACTCAACTGGTTGTTT 3’- reverse primer for generating ANLN (1-91aa); ANLN (57-91aa) rev_ anln _151 CTCTGCAAGTTTTTGCATAC 3’- reverse primer for generating ANLN (1-151aa); ANLN (22-151aa); ANLN (57-151aa); ANLN (91-151aa) rev_ anln _450 CTTTTCATTCTTTTCTGCACTCC 3’- reverse primer for generating ATAA ANLN (91-450aa) rev_ anln _300 AGTAGATTTCACTGGAGAAG 3’- reverse primer for generating ANLN (1-300aa)
5’_mDia2 ATGGAACGGCACCAGCCG 5’-initial forward primer of mDia2
117 Supplementary
3’_mDia2 TTATAAAGCTCGTAATCTTGCC 3’-end reverse primer of mDia2 fwd_mDia2_89 CCACTTCCCAACCTGAAG 5’- forward primer for generating
NT89-533 fwd_mDia2_561 -CCTTTGCCTCCCTCTAAAGA 5’- forward primer for generating CT rev_ mDia2_533 TTTTTTCTGCAATTCAGCCTGA 3’- reverse primer for generating
GT NT89-533
5’_importin β2 ATGGTGTGGGACCGGCAAACC 5’-initial forward primer of importin β2 AAGAT
3’_importin β2 AATTTGTGGTATTTTTCGACGT 3’-end reverse primer of importin β2 TCTGCGAG
118 Supplementary
Table 6.3 Oligonucleotides used to generate mutant constructs
Forward primer (5’!3’) Reverse primer (5’!3’) Feature (paired with rev_ anln _151) (paired with 5’_anln) TCTTGTACAAAACCATCGCC AGAAACTTCTACTTCAGTG Oligos for generating the ATCAGCAGCAGCATGTTCTG TTGTCAGAACAACGACGAC ANLN (1-151aa)-KKR ACAACACTG GTGATGGCGATGGTTTTGT (68-70)-AAA mutant ACAAGATTT GACAACACTGAAGTAGAAG AGATTTTGCAGATGTCGAC Oligos for generating the TTTCTAACTTGGCAAATAAA TCAACTGGTTGTTTATTAC ANLN (1-151aa)-E83A CAACCAGTTGAGTCGACATC GCAAGTTAGAAACTTCTAC mutant T TTCAGT AACACTGAAGTAGAAGTTTC ACAAGATTTTGCAGATGTC Oligos for generating the TAACTTGGAAGCAAAACAA GACTCAACTGGTTGTTTAC ANLN (1-151aa)-N84A CCAGTTGAGTCGACATCTGC GTTCCAAGTTAGAAACTTC mutant A TACTTC ACTGAAGTAGAAGTTTCTAA TGGAGAACAAGATTTTGCA Oligos for generating the CTTGGAAAATAAAGCACCA GATGTCGACTCAACTGGAC ANLN (1-151aa)-Q86A GTTGAGTCGACATCTGCAAA GTTTATTTTCCAAGTTAGA mutant A AACTTCTAC Forward primer (5’!3’) Reverse primer (5’!3’) Feature (paired with 3’_anln_Lic) (paired with 5’_anln_Lic) TCTTGTACAAAACCATCGCC AGAAACTTCTACTTCAGTG Oligos for generating the ATCAGCAGCAGCATGTTCTG TTGTCAGAACAACGACGAC ANLN (FL)-KKR ACAACACTG GTGATGGCGATGGTTTTGT (68-70)-AAA mutant ACAAGATTT Forward primer (5’!3’) Reverse primer (5’!3’) Feature (paired with rev_ anln _91) (paired with 5’_anln) CCAAGGTCTATGACTCATGC TTCTGAAAGTGGCTGTCTA Oligos for generating the TGCAGCAGCTAGACAGCCA GCTGCTGCAGCATGAGTCA ANLN (1-91aa)-KR T (38-39)-AA mutant
ATGACTCATGCTAAGCGAGC TGCTTCTGAAAGTGGCTGT Oligos for generating the TGCACAGCCACTTTCAGAA GCAGCTCGCTTAGCATGAG ANLN (1-91aa)-R41A T mutant
ACTCATGCTAAGCGAGCTAG GTTACTTGCTTCTGAAAGT Oligos for generating the AGCACCACTTTCAGAAGCA GGTGCTCTAGCTCGCTTAG ANLN (1-91aa)-Q42A A CATG mutant
CATGCTAAGCGAGCTAGAC CTGCTGGTTACTTGCTTCTG Oligos for generating the
119 Supplementary
AGGCAGCATCAGAAGCAAG ATGCTGCCTGTCTAGCTCG ANLN (1-91aa)-PL TAAC CT (43-44)-AA mutant
TCTATGACTCATGCTAAGCG GGGCTGCTGGTTACTTGCT Oligos for generating the AGCTGCAGCAGCAGCATCA TCTGATGCTGCTGCTGCAG ANLN (1-91aa)-RQPL GAAGCAAGTAAC CTCGCTTAGCATGAGT (41-44)-AAAA mutant
120