Functional Characterisation of Kif18A in Xenopus laevis and Generation of Inhibitor-Sensitive Kinesin Chimeras

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Functional Characterisation of Kif18A in Xenopus laevis and

Generation of Inhibitor-Sensitive Kinesin Chimeras

Dissertation submitted for the degree of Doctor of Natural Sciences (Dr.rer.nat.)

Presented by

Martin Michael Möckel

at the

Faculty of Sciences Department of Biology

Date of oral examination: 13

^th

of November 2015 First referee: Prof. Dr. Thomas U. Mayer Second referee: Prof. Dr. Martin Scheffner Third referee: Prof. Dr. Stefan Westermann

Konstanzer Online-Publikations-System (KOPS) URL: http://nbn-resolving.de/urn:nbn:de:bsz:352-0-310495

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In vitro veritas.

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1 Summary

Kinesin motor proteins guide a multiplicity of cellular processes including transport of cellular components along microtubules, ciliogenesis and spindle formation in M-phase.

During the work of this thesis an approach that enables targeting of kinesins with a well characterised, cell-permeable small molecule inhibitor of the human kinesin Eg5, S-trityl cysteine (STLC), has been established. The compound specifically inhibits Eg5 through binding to an allosteric site within the motor domain, blocking ADP release and hence the enzyme´s ATPase cycle. It could be shown that the transfer of this allosteric binding site to other kinesins renders them sensitive towards STLC, but does not drastically affect their native activity in vitro. This domain-swap approach could be a valuable tool in the future to target kinesins for which no potent small molecule inhibitors are available yet with high spatio-temporal precision.

Furthermore, the M-phasic kinesin-8 Kif18A could be identified in the African clawed frog Xenopus laevis. It could be shown that the kinesin is a plus-end-directed, processive motor with microtubule bundling activity. Additionally, domains important for Kif18A´s function were identified. The kinesin becomes abundant during meiotic resumption and is expressed throughout early embryonic divisions in Xenopus. It could be shown that Kif18A is required for faithful spindle assembly in M-phasic Xenopus egg extract. Furthermore, the motor protein is able to complement the activity of its human homologue in human cultured cells, suggesting a conserved function for Kif18A in higher vertebrates. Notably, preliminary data indicate an essential role of the kinesin during the early embryonic cell divisions of Xenopus laevis, highlighting the function and importance of kinesins in development of higher vertebrates.

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Summary/Zusammenfassung

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1 Zusammenfassung

Kinesin Motorproteine steuern eine Vielzahl zellulärer Prozesse, wie zum Beispiel den Transport zellulärer Komponenten, Ziliogenese oder den Aufbau der mitotischen Spindel.

Im Zuge dieser Arbeit wurde eine Methode entwickelt, die es ermöglicht, verschiedene Kinesine mit einem Inhibitor des Motorproteins Eg5 zu inaktivieren. Bei dem Inhibitor handelt es sich um die zellpermeable, organische Verbindung S-trityl Cystein (STLC). STLC bindet spezifisch an eine allosterische Bindestelle in der Eg5 Motordomäne, inhibiert das Freiwerden von ADP vom Enzym und somit den ATPase Zyklus des Motors. Der Transfer der allosterischen Bindestelle von Eg5 in andere Kinesine führte zur Inaktivierung dieser durch STLC, ohne jedoch die generelle Enzymaktivität in Abwesenheit von STLC stark zu beeinflussen. Der beschriebene Transfer der allosterischen Bindestelle könnte in genetisch veränderten Zellen benutzt werden, um Kinesine, für die es noch keine spezifischen Inhibitoren gibt, zeitlich präzise zu inaktivieren.

Weiterhin konnte das Kinesin-8 Kif18A im afrikanischen Krallenfrosch Xenopus laevis identifiziert werden. Es konnte gezeigt werden, dass das Kinesin weite Strecken an Mikrotubuli zu deren Plus-Enden zurücklegt und Mikrotubuli bündeln kann. Weiterhin wurden Proteindomänen, die für die Funktion des Kinesins wichtig sind, identifiziert. Kif18A ist präsent in Meiose und in den frühen embryonalen Teilungen des Klauenfrosches. Das Kinesin wird für den Aufbau der Spindel in M-Phase Eiextrakt benötigt und kann die Funktion des humanen Homologs in menschlichen, kultivierten Zellen übernehmen. Vorläufige Experimente weisen auf eine essentielle Funktion von Kif18A in den frühen embryonalen Zellteilungen von Xenopus laevis hin. Dies unterstreicht die Bedeutung von Kinesinen während der Embryonalentwicklung höherer Wirbeltiere.

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Introduction

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2 Introduction

One fundamental principle in nature is the proliferation of living cells by cycles of growth and subsequent division. This phenomenon applies for any organism from single celled prokaryotes to multicellular eukaryotes and ensures the propagation of their genetic material, a feature termed genetic fitness.

In eukaryotes, there generally exist two ways to propagate the genome: Equal distribution between daughter cells is achieved during equational divisions (mitosis), a prerequisite e.g. for the development and growth of a multicellular organism. In contrast to mitosis, meiosis consists of two consecutive divisions: a reductional division (meiosis I), where the typically diploid set of chromosomes is “reduced” to a haploid one, followed by an equational division where the remaining sister chromatids are separated from each other (meiosis II).

2.1. Principles of the eukaryotic cell cycle

The eukaryotic cell cycle represents the order of events that are required for cell growth and subsequent division. Genome replication during S (synthesis) phase and its division during M (mitotic) phase are typically separated by two G (gap) phases in which cells increase their biomass. Keeping the strict order of these four phases is a prerequisite for functional cell divisions and genetic stability in eukaryotes. Therefore, transitions between the different cell cycle stages have to be unidirectional, switch-like and controlled by check points that block cell cycle progression if specific requirements are not met (Morgan, 2007).

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Introduction

12 Figure 2.1. Schematic for the eukaryotic cell cycle. The eukaryotic cell cycle is divided into four phases.

Chromosome duplication during S-phase and their separation during M-phase are divided by two Gap-phases (G1 and G2) where cells increase their biomass. Cell cycle transitions are switch-like and controlled by checkpoints, like the DNA damage checkpoint during G2-phase and the spindle assembly checkpoint during M-phase. Illustration adapted from Morgan, 2007.

At the heart of cell cycle regulation lie the oscillating activities of master kinases, the Cyclin- dependent kinases (Cdks), and their antagonising phosphatases. These key regulators itself are controlled by the activity of E3 ubiquitin ligases and subsequent proteasome-mediated proteolysis. There exists extensive crosstalk between kinases, phosphatases and E3-ligases to ensure the right order and timing of events during the cell cycle.

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Introduction

13 2.2. Mitotic progression

During mitosis the duplicated chromosomes get divided into the newly forming daughter cells.

Mitosis itself consists of different subphases that constitute the order of events as mitosis progresses and can be distinguished on visual and molecular level (Pines and Rieder, 2001).

Figure 2.2. The different mitotic phases visualised by fluorescence microscopy. Mitosis can be divided into five different subphases: prophase (A, B), prometaphase (C, D), metaphase (E), anaphase (F) and telophase/cytokinesis (G, H). In mitosis chromosomes are separated into the newly forming daughter cells during anaphase, whereas the cytoplasm is divided later, during cytokinesis. Pictures obtained from Rieder and Khodjakov, 2003.

The master regulator of mitotic entry is the Cyclin-dependent kinase 1 (Cdk1) in complex with its co-activator Cyclin B. Cdk1 activity is regulated by complex networks, resulting in a bi-stable, switch-like activation/inactivation (Ferrell, 2013). Upon full Cdk1 activation mitotic entry is triggered. During prophase, chromosomes condense and the nuclear envelope breaks down,

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Introduction

14 which is facilitated by the Cdk1-mediated phosphorylation of lamins and nuclear pore complexes (Peter et al., 1990). The breakdown of the nuclear envelope makes chromosomes accessible to their separating machinery, the mitotic spindle (see also paragraph 2.4. “The mitotic spindle”). Prior to nuclear envelope breakdown the two spindle poles, which consist of centrosomes in animals, are separated. This is a crucial process required for spindle assembly and it depends on the motor protein Eg5 (Kapitein et al., 2005; Sawin et al., 1992; Fig.2.2.B; see paragraph 2.6. for details on the kinesin Eg5). In prometaphase kinetochores of chromosomes are attached to microtubules emanating from opposite spindle poles (Fig.2.2.C). In contrast to interphase microtubules, these mitotic microtubules are growing and shrinking much faster in order to rapidly roam the cytoplasm in a search and capture for kinetochores (Rieder and Khodjakov, 2003; the underlying mechanism of rapid microtubule growth and shrinkage is explained in paragraph 2.3.). Kinetochores are first laterally attached to microtubules and then migrate towards the middle of the spindle, a process that is termed congression (Rieder and Salmon, 1994; Fig.2.2.D). Here, lateral attachments are transformed to end-on attachments, meaning that microtubules are connected to kinetochores with a 90 degree angle to the chromosome axis. Chromosome congression, as well as the formation of stable end-on attachments, depends on chromosome transport to the microtubule plus tip by CENP-E and the termination of existing lateral attachments by the microtubule depolymerase MCAK (Shrestha and Draviam, 2013). After end-on attachment is achieved, microtubule-kinetochore interactions are stabilised, resulting in the formation of microtubule bundles connected to kinetochores and consisting of up to 28 individual microtubules (k-fibres) in higher eukaryotes (McEwen et al., 1997). Stably attached chromosomes then undergo back and forth movements with abrupt switches in their direction along the spindle pole-to-pole axis (Jaqaman et al., 2010; Skibbens et

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Introduction

15 al., 1993). These oscillations are thought to be a quality control mechanism for stable, correct end-on attachment of sister kinetochores to k-fibres from opposite spindle poles (amphitelic attachment). Before the onset of anaphase, oscillatory movements are suppressed by the activity of the kinesin Kif18A (see paragraph 2.7. for details on Kif18A and further speculation about the requirements of chromosome oscillations), resulting in the tight alignment of chromosomes in the metaphase plate (Fig.2.2.E). Only when all sister kinetochores are attached in an amphitelic fashion, the spindle assembly checkpoint is satisfied, leading to a release of inhibition of the E3 ubiquitin ligase APC/C (anaphase promoting complex/cyclosome; King et al., 1995; Musacchio, 2011). The APC/C then ubiquitinates the Cdk1 co-activator Cyclin B (Glotzer et al., 1991) and the separase inhibitor securin (Yamamoto et al., 1996). This leads to the proteasome-dependent destruction of these proteins, resulting in the activation of separase (Ciosk et al., 1998) and a switch-like drop in Cdk1 activity. Following Cdk1 inactivation, Cdk1- dependent inhibitory phosphorylations on mitotic phosphatases are removed, leading to their activation and hence a drop in phosphorylation levels of mitotic kinase substrates (Grallert et al., 2015; Wu et al., 2009). The protease separase cleaves the cohesion ring, a protein complex that topologically entraps DNA from two sister chromatids (Gruber et al., 2003; Uhlmann et al., 2000). Cohesin cleavage results in the loss of cohesion between sister chromatids. The chromatids are then rapidly separated to opposing spindle poles by the shrinkage of k-fibres in anaphase, which completes an important step in cell division, karyokinesis (Fig.2.2.F). After chromatid separation, stable Ran-dependent, antiparallel microtubule bundles form between the segregating chromosome masses (Walczak and Heald, 2008). These antiparallel bundles, which have their plus ends pointing inside the overlap, are termed the central spindle (Fig.2.2.G). The central spindle determines the position of the cleavage furrow by acting as a

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Introduction

16 platform for microtubule-interacting proteins (MAPs) and molecular motors that transport important cytokinesis factors to the centre of the dividing cell (Glotzer, 2009). A contractile acto-myosin II ring is established and activated through kinesin-dependent transport of GEFs and GAPs of the small G protein RhoA to the centre of the midbody. Full cycling of RhoA through its GTPase cycle leads to the ingression of the cleavage furrow (Glotzer, 2005; Miller and Bement, 2009). After furrow ingression, cytokinesis is completed and only the midbody, composed of the compacted spindle midzone, remains as a physical link between the two newly formed daughter cells (Fig.2.2.H). During abscission, this last connection is finally terminated and the midbody remnant is internalised by one of the two daughter cells.

2.3. Microtubules are at the core of the mitotic spindle

Microtubules are polar, tube-like structures built of repetitive α-β-tubulin heterodimers. Their dynamic nature makes these non-equilibrium polymers ideal to fulfil the role as molecular scaffold that drives chromosome segregation during M-phase. In the following paragraph, the molecular basis of tubulin and microtubules that result in their special polymer behaviour will be explained.

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Introduction

17 Figure 2.3.1. Schematic of a microtubule and structure of the α-β-tubulin dimer. (A) Illustration of a polar microtubule consisting of repetitively arranged α-β-tubulin dimers within 13 parallel protofilaments that interact with each other laterally. Shifted lateral interaction between protofilament 1 and 13 gives rise to the microtubule seam. (B) Crystal structure of a polar α-β-tubulin dimer within a microtubule protofilament (β-tubulin in blue and α-tubulin in green). The nucleotide (GTP or GDP, orange) in the nucleotide binding site of β-tubulin is exchangeable, whereas the binding site of α-tubulin is buried within the stable α-β-dimer interface and the nucleotide is therefore not exchangeable. Structure from Alushin et al., 2014.

Within a microtubule, α-β-tubulin heterodimers are associated with each other longitudinally, forming repetitive polar protofilaments. In addition, protofilaments are attached laterally to each other, leading to the formation of the tube-like microtubule. In vivo most microtubules consist of 13 protofilaments. Since the longitudinal contact sites between tubulin subunits of different protofilaments are shifted by approximately 0.9 nm, α-β-tubulin dimers follow a left- handed helical pathway around the microtubule between the different protofilaments, with the individual protofilaments running completely parallel to each other. This results in the formation of a mismatch between protofilaments 13 and 1, the microtubule seam. Here, the lateral contact is made between α- and β-tubulin, instead of an α-α or β-β interaction, leading to a weaker association of these two protofilaments within the microtubule (Nogales, 2000;

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Introduction

18 Wade, 2009; Fig.2.3.1.A). This might have implications during microtubule growth, where an open, sheet-like structure could be formed first which might then be closed at the seam region as polymerisation advances (see Fig.2.3.2. “polymerisation” for a possible model).

α- and β-tubulin are mutual-activating GTPases that are conserved over 50 % on sequence level

and display almost identical structure. The two homologues form a polar heterodimer in solution. The GTP binding site of tubulin is located at its N-terminus that points towards the longitudinal interaction surface (Fig.2.3.1.B). The nucleotide bound to α-tubulin is buried within the α-β-interaction surface called the N-site (non-exchangeable), resulting in a tight interaction between α- and β-tubulin within a heterodimer and hence the inability of Pi release from α- tubulin. In contrast to α-tubulin, the nucleotide-binding site of β-tubulin is accessible (E-site, exchangeable) and the binding of another α-subunit to this GTP-bound β-tubulin mediates nucleotide hydrolysis and subsequent Pi release, which bends the two associated monomers, and results in a slightly curved heterodimer (Nogales, 2000).

In all eukaryotic cells a scaffold containing the tubulin homologue γ-tubulin caps the less dynamic, α-terminal minus end of microtubules and thereby restricts growth to the β-terminal plus end. Here, the binding of the α-subunit from a free tubulin dimer to a GTP-bound β-subunit at the plus end of a microtubule mediates GTP hydrolysis and Pi release. Therefore β-tubulin subunits in the lattice of a microtubule, distant to the plus tip, are in a GDP-bound state, whereas plus tip located β-tubulins form the so-called GTP cap. Since the GDP state favours a slightly curved, non-linear conformation within heterodimers, friction forces between laterally attached protofilaments are generated (Nogales, 2000). However, these forces can be balanced out by the dimers in the GTP cap region, because they form tight lateral interactions and

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Introduction

19 thereby stabilise the microtubule lattice in a straight conformation. If the GTP cap is lost due to hydrolysis rates exceeding the incorporation of new dimers (mediated by associated proteins or stochastic events), the dimers at the microtubule tip adopt a bent conformation and the α- tubulin is rotating slightly, leading to a weakening of lateral attachments at the tip (Zhang et al., 2015). Consequently, the frictional forces inside the lattice are no longer balanced out and protofilaments start to peel apart, resulting in the rapid shrinkage and depolymerisation of the microtubule (Desai and Mitchison, 1997). This non-equilibrium behaviour of microtubules can be summarised in the model of dynamic instability (Fig.2.3.2.).

Figure 2.3.2. Model for dynamic instability of microtubules. During polymerisation α-β-tubulin dimers that contain GTP at the E-site are attached to a growing microtubule plus tip. The tube is closed by lateral interactions between protofilaments and the final zippering between protofilament 1 and 13 at the microtubule seam region.

The switch from polymerisation to depolymerisation is termed catastrophe. Microtubules depolymerise when the cap of GTP-bound α-β-tubulin dimers at the plus tip is lost and hence the friction forces within the GDP-α-β- tubulin-containing microtubule lattice are no longer balanced. Depolymerisation is mediated by peeling of the protofilaments. The switch from depolymerisation back to polymerisation is termed rescue. Model adapted from Walczak et al., 2010.

Dynamic instability is described by four parameters: rates of microtubule growth and shrinkage and the transition frequencies from growth to shrinkage (catastrophe) and shrinkage to growth (rescue; Mitchison and Kirschner, 1984). These parameters depend on the molecular properties of tubulin, briefly mentioned above, and are an inherent property of all microtubule-based processes, as for example inside the mitotic spindle.

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Introduction

20 2.4. The mitotic spindle

The mitotic spindle serves as the chromosome-segregating machinery during cell division. Its core structure is composed of microtubules that are bundled at their minus ends by two major microtubule organising centres, the centrosomes, which are lacking in some special forms of cell divisions (e.g. female meiosis in vertebrates or generally in plants).

Figure 2.4. Schematic of the mitotic spindle during metaphase in vertebrates. Microtubules of the mitotic spindle are bundled at their minus ends by the centrosome that contains many microtubule organising centres (MToCs).

Microtubules within the mitotic spindle are divided into different subclasses. Astral microtubules make contact to the cell cortex. Interpolar microtubules are directed towards the cell middle and span over the chromosome masses while being connected to interpolar microtubules from the opposite pole. Kinetochore-attached k-fibre microtubules help arranging chromosomes inside the mitotic spindle and are required for their segregation during anaphase. At branching points, augmin-dependent recruitment of MToCs to the spindle leads to microtubule branching and increase of the microtubule mass. Illustration adopted from Walczak et al., 2010.

The microtubules inside the spindle can be separated into different subclasses (Walczak and Heald, 2008): Astral microtubules are pointing towards the cell periphery, contacting the cell cortex and are important to position and anchor the spindle inside the cell. Interpolar microtubules span over the chromosomes towards the opposite spindle pole and are connected to interpolar microtubules from the other pole. Here, pushing and pulling forces exerted by

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Introduction

21 motors like cytoplasmic dynein and the tetrameric kinesin Eg5 that crosslink these interpolar microtubules are required to establish and maintain the proper pole-to-pole distance (Tanenbaum et al., 2009). However, it is likely that dynein does not directly counteract Eg5 in a simple push-pull model at the same microtubule overlap and more players are probably involved in order to achieve the required pole-to-pole distance during mitosis (Florian and Mayer, 2012). Kinetochore-attached microtubules (k-fibres) are microtubule bundles that consist of up to 28 individual microtubules in vertebrates. K-fibres are the microtubules that are physically connected to chromosomes and hence have a major impact on chromosome movements during mitosis. For example they are directly involved in sister chromatid separation during anaphase by exerting opposing forces on sister kinetochores that lost their cohesion.

Microtubule nucleation is driven by at least three different sources inside the mitotic spindle. At the centrosome, γ-tubulin ring complexes (γ-TuRCs) are concentrated and guide microtubule nucleation. Thirteen γ-tubulins are arranged with the help of additional proteins, the γ-tubulin complex proteins (GCPs), to form the γ-TuRC, which acts as a nucleation source and capping factor at microtubule minus ends. By interacting with the terminal α-tubulins of a microtubule, they give them their typical 13-protofilament structure in cells (Kollman et al., 2011). The γ-TuRC forms the minimal basis of all microtubule organising centres (MToCs) in both centrosomal and acentrosomal spindles (Walczak and Heald, 2008). At the microtubule lattice, augmin complexes can recruit additional γ-TuRCs, resulting in the nucleation of new microtubules from branching points (Goshima et al., 2008; Petry et al., 2013). Strong Ran-dependent microtubule nucleation can be observed around chromosome arms (Gruss et al., 2001). It is likely that the Ran- dependent pathway accounts for the majority of microtubule nucleation within the spindle.

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Introduction

22 When polymerising near chromosomes, these microtubules are pushing their minus ends towards the spindle poles and can then be caught and centred by MToCs. This type of nucleation is believed to also account for k-fibre formation. Here, microtubules could nucleate directly at the kinetochore and therefore increase the likelihood of chromosomes to be captured by microtubules inside the mitotic spindle during the early stages of mitosis (Walczak and Heald, 2008).

During mitotic progression the spindle is changing its shape and properties in order to faithfully perform the task of karyokinesis, as already described briefly in paragraph 2.2. “Mitotic progression”. These drastic changes within the spindle are brought about by microtubule associated proteins (MAPs) and motor proteins like kinesins and cytoplasmic dynein. These spindle-associated proteins modulate the intrinsic properties of microtubule dynamic instability and exert forces on the spindle itself, as well as on chromosomes and the cell cortex throughout mitosis. Therefore the function and regulation of these proteins is crucial for successful cell division and have been subject of extensive studies throughout the past decades (Gadde and Heald, 2004). In the next paragraph, kinesin motor proteins and the function and molecular properties of two mitotic kinesins, Eg5 and Kif18A, which were subject of this study, will be introduced in further detail.

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Introduction

23 2.5. The kinesin superfamily of proteins

Kinesins are motor proteins that convert chemical energy into mechanical force, resulting in their movement along microtubules. These molecular machines most often form dimeric or tetrameric structures to generate movement through cooperation of two or more of their globular motor domains, a structural feature shared by all kinesins (Vale and Fletterick, 1997;

see Fig. 2.5.1.A). The motor domain can undergo structural rearrangements during its ATP hydrolysis cycle and interacts with microtubules, as described in more detail in the next paragraphs. During evolution, the feature of interaction with microtubules and (most often) motility along microtubules has been utilised to fulfil a variety of functions inside cells. The distinct requirements of a kinesin are tailored through fine-tuning of the mechanochemical properties of the motor domain, as well as by the special characteristics of the less conserved non-motor elements and sometimes also associated factors (see Fig. 2.5.1.A for a schematic depiction of a dimeric kinesin). This resulted in the establishment of more than 40 different kinesin genes in humans. These different kinesins can be clustered into 14 subfamilies according to their degree of conservation in both the motor domain and the less conserved non-motor parts (Lawrence et al., 2004; Miki et al., 2005). Kinesins can also be clustered depending on the position of their motor domain (N-terminal, C-terminal or internal) and their directionality of movement to either the plus or minus end of microtubules (Vale and Fletterick, 1997). The work of this thesis focused on plus end-directed mitotic kinesins of the kinesin-5 and -8 families that harbour an N-terminal motor domain. Therefore the following two paragraphs will describe the general mode of action and underlying molecular properties based on structures of N-terminal plus end-directed motors (kinesin-1 and kinesin-5), before details on both Eg5 (kinesin-5 family) and Kif18A (kinesin-8 family) are outlined.

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Introduction

24 2.5.1. Structure of the kinesin motor domain

Figure 2.5.1. Schematic of a dimeric kinesin and structure of the hsEg5 motor domain. (A) Schematic of a dimeric N-type kinesin. The different domains are labelled on the left side and their functions within the kinesin are briefly described on the right. (B) Structure of the Eg5 motor domain from Homo sapiens (hs) obtained by crystallography by Parke et al., 2010 (PDB ID: 3HQD). Important elements are colour-coded: ATP-binding region (switch I, II and p- loop) in blue, bound ATP-analogue in red, transducer helix 4 in green, microtubule-binding loops 8 and 12 in dark green, helix 6 in orange and neck linker region in red.

The core of the kinesin motor domain is formed by eight mostly parallel β-strands (Fig.2.5.1.B, cyan) that are covered by three α-helices to each side. The ATP binding pocket located towards the distal tip of the molecule is relatively open and formed by four loops that emerge from the C-termini of four core beta strands. Three of these strands form the p-loop and the switch I and II elements (Fig.2.5.1.B, blue), which are coordinating nucleotide binding and hydrolysis and undergo conformational changes during the hydrolysis cycle as explained in the next paragraph.

The arrangement of these loop elements around the nucleotide is similar in the myosin family and small G-proteins and although poorly conserved in sequence, all three classes of enzymes share a similar molecular mode of action during their nucleotide hydrolysis cycle (Vale, 1996).

In the depicted structure, the microtubule interaction surface lies on the bottom of the molecule. The motor domain interacts with both α- and β-tubulin of a tubulin heterodimer.

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Introduction

25 Interactions are mediated, among others, by kinesin´s loop 8 and loop 12 towards β-tubulin (Fig.2.5.1.B, dark green) and by kinesin´s helix 4 towards both α- and β-tubulin (Fig.2.5.1.B, light green). Helix 4 is also termed the relay helix, as it changes its position and angle within the motor domain depending on the nucleotide state and has therefore once been compared to a piston inside an Otto-engine (Vale and Milligan, 2000). At the C-terminal end of the motor domain, distal to helix 6, lies another important mechanical element, a flexible stretch called the neck linker (Fig.2.5.1.B, red). If helix 4 is the piston, then the neck linker can be viewed as the crankshaft inside a kinesin motor. The neck linker can interact with the motor domain in a nucleotide-dependent manner (see paragraph 2.5.2.). It transduces the force generated through ATP hydrolysis from one motor domain to the other and is involved in shifting/gating the ATPase cycles of both motor heads (described in paragraph 2.5.2.). Since the neck linker is very flexible it also ensures that both heads in a kinesin dimer have a high degree of rotational freedom when moving along a microtubule (Vale and Milligan, 2000).

2.5.2. The mechanochemical cycle of plus end-directed dimeric kinesins

The unidirectional stepping of kinesin along microtubules is achieved by alternating cycles of microtubule binding and unbinding, coupled to ATP hydrolysis. These processes have to be coordinated between the two motor domains by gating mechanisms in order to achieve a hand- over-hand movement of both domains. Recent co-crystal structures of kinesin bound to an α-β- tubulin dimer in both ATP-bound and nucleotide free form shed light into most of the structural arrangements and changes during kinesin´s ATPase cycle, some of which were still debated after 30 years of kinesin research (Cao et al., 2014; Gigant et al., 2013; Wang et al., 2015a).

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Introduction

26 Figure 2.5.2. Model for the kinesin mechanochemical cycle. (A-F) Illustration depicting a simplified version of the mechanochemical cycle of a kinesin dimer that encounters a microtubule and then walks along it. Details are described in the figure and in the text. The two heads (1 and 2) are colour-coded and the nucleotide state of the heads is abbreviated as T (ATP bound), D (ADP bound) and - (apo form, no nucleotide bound). (G, H) Different zooms into the co-crystal structure of kinesin-1 motor domain (green) bound to a tubulin dimer (blue; Gigant et al., 2013). (G) Focus on the N-terminal part of helix 4 that elongates upon tubulin binding and is depicted in darker green. (H) Rearrangement of helix 6 and the p-loop upon ATP binding (green, vs. ADP bound state in yellow) that leads to aromatic stacking between F318 of helix 6 and Y84 of the p-loop. (I) Comparison of the arrangement of helices 4 and 6 between the neck linker-docked and -undocked state within the motor domain of kinesin-1 (Wang et al., 2015a).

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Introduction

27 When a kinesin dimer encounters a microtubule, only one of its two heads can bind to the protofilament in the nucleotide-free apo form, because the second head is positioned too far away from the tubulin binding interface (Vale and Milligan, 2000; Fig.2.5.2.A). Upon microtubule binding, helix 4 interacts with α-tubulin, resulting in the elongation of the helix´ N-terminus and its translocation by several angstroms (Gigant et al., 2013; Fig.2.5.2.G). This allows reordering of the switch regions to favour an ATP-bound state (Fig.2.5.2.B). As a consequence, the p-loop is rotated into a position where aromatic stacking between a side chain in this loop and the more than 250 amino acids apart C-terminal helix 6 can occur (Gigant et al., 2013; Fig.2.5.2.H). The resulting parallel alignment of these two elements pulls on helix 6 so that it loses its last winding to the C-terminus (Fig.2.5.2.I). This partial unwinding of helix 6 and the rearrangement of helix 4, as well as the p-loop rotation described above, create a cavity suitable for stable neck linker docking onto head 1 (Fig.2.5.2.C and I). Neck linker binding leads to tension on the rear head towards the front. This tension is released by force transduction over the coiled-coil dimer interface that results in a rearward-to-forward swing of the second head into the direction of the microtubule plus tip (Vale and Milligan, 2000; Fig.2.5.2.C). Since the second head is now adjacent to the next α-β-tubulin binding interface, it can bind to it in the nucleotide-free apo state similar to head 1 before (Fig.2.5.2.C). The neck linker bound to head 1 that now makes contact with the catalytic core of the motor domain stimulates its ATPase activity (Wang et al., 2015a). Here, a water molecule adjacent to the γ-phosphate of the bound ATP is activated by a conserved glutamate in switch II leading to ATP hydrolysis by base catalysis (Gigant et al., 2013;

Fig.2.5.2.C). After hydrolysis the free γ-phosphate is quickly released from the motor.

Subsequently the switch regions, as well as the p-loop, adopt a conformation that lead to a shortening and translocation of helix 4 which results in a reduced affinity of the helix towards α-

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Introduction

28 tubulin. Because the p-loop rotates back, the pull on helix 6 is released, leading to its re- elongation and the closure of the neck linker-docking site followed by the neck linker´s dissociation from the motor domain 1 (Wang et al., 2015a; Fig.2.5.2.D). Head 2 that is now bound to the tubulin dimer in front of head 1 can meanwhile bind ATP in a similar fashion as described above for head 1 (Fig.2.5.2.D). Subsequent neck linker-docking leads to the rearward- to-forward swing of the first, now weakly associated, head into the direction of the microtubule plus tip (Fig.2.5.2.E). Experiments with external load application on neck linker-containing kinesin motors suggest that the increase of microtubule minus end-directed tension on a front- tethered motor domain (here on head 1) reduces its affinity for nucleotide (Uemura and Ishiwata, 2003). It is probably this increase of minus-end directed tension on head 1 that allows it to release the ADP molecule (Fig.2.5.2.F). The resulting nucleotide-free apo form has again high microtubule affinity and can tightly bind to the proximate α-β-tubulin dimer (Fig.2.5.2.F), initiating the next repetition of the described mechanochemical cycle that represents the basis of processive kinesin movement along microtubules (iteration of cycles from Fig.2.5.2.D to E to F).

2.6. The kinesin-5 Eg5 and its allosteric inhibitors targeting loop 5

As a member of the kinesin-5/BimC family (Miki et al., 2005), Eg5 forms a homo-tetrameric structure through parallel coiled-coil interactions within each dimer and antiparallel coiled-coil interactions between the two dimers within a bipolar assembly domain (Scholey et al., 2014).

These extensive parallel and antiparallel coiled-coil interactions ensure a structural arrangement where two N-terminal motor domains are located on each side of the rod-like tetramer (Fig.

2.5.3.A). Eg5 plays an important role in bipolar spindle assembly during mitosis (Sawin et al.,

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Introduction

29 1992; Walczak et al., 1998), as it can crosslink antiparallel microtubules emanating from opposing spindle poles. Eg5 pushes these antiparallel microtubules apart by plus tip-directed forces, leading to the separation of the spindle poles and the establishment of a bipolar spindle (Kapitein et al., 2005; see also 2.2. “Mitotic progression” and 2.4. “The mitotic spindle”).

Figure 2.6. Model for Eg5´s mode of action and structure of the Eg5 motor domain together with the allosteric inhibitor S-trityl cysteine (STLC). (A) Eg5 tetramers crosslink interpolar microtubules emanating from opposite spindle poles. Due to directional motor activity on both sides of the rod-like molecule, the antiparallel microtubules can be slid apart. This figure was directly extracted from (Scholey et al., 2014). (B) Crystal structure of the human Eg5 motor domain in complex with ADP and its inhibitor S-trityl cysteine (STLC; Kaan et al., 2010; PDB ID: 2WOG).

(C) Zoom into the structure from (B) to illustrate the relevant side chains of residues (red) involved in STLC (yellow) binding. Residues were selected according to Liu et al., 2011. Secondary structures that harbour the coloured residues are labelled.

In a phenotype-based screen for small molecule inhibitors that block mitotic progression without targeting tubulin itself, a compound that specifically inhibits the motor activity of Eg5 was discovered. Since chemical inhibition of Eg5 leads to monopolar spindles with their two spindle poles collapsed in the middle of the mitotic cell, the compound was termed Monastrol (Mayer et al., 1999). Monastrol-treated cells are arrested in mitosis due to mal-attached, mono- oriented kinetochores that activate the spindle assembly checkpoint (Kapoor et al., 2000). The

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30 binding region of Monastrol within Eg5 is mainly loop 5 that bridges the N- and C-terminal part of helix 2. In the past decade, several loop 5-targeting compounds with different lead structures were discovered (reviewed in Wojcik et al., 2013). One of them, S-trityl cysteine (STLC), is a cell- permeable inhibitor with low micromolar IC50 values both in vitro and in vivo (Skoufias et al., 2006). Eg5 bound to STLC is locked in an ADP-bound transition state with low affinity for microtubules and loop 5 is “cradling” the compound, adopting a closed conformation (Kaan et al., 2010; Fig.2.6.B). In addition to its interaction with loop 5, STLC is “sandwiched” by two alpha helices (α2 and α3). The residues important for STLC interaction have been identified from structural data (Liu et al., 2011) and are almost exclusively located in loop 5 and the alpha helices 2 and 3 (Fig. 2.6.C).

2.7. The kinesin-8 family with focus on its member Kif18A

Members of the kinesin-8 family are N-terminal, dimeric kinesins (Miki et al., 2005; see Fig.2.7.C schematic of Kif18A dimer). At least three of them, Kif18A, Kif18B and Kif19A, are regulating microtubule dynamics/length in various different cellular processes (Mayr et al., 2007; Niwa et al., 2012; Stout et al., 2011; Tanenbaum et al., 2011).

Kif18A and its homologues (Kip3 in Saccharomyces cerevisiae, heterodimeric Klp5/6 in Schizosaccharomyces pombe and Klp67A in Drosophila melanogaster) are M-phase kinesins that regulate microtubule dynamics during cell division. The structure of the human Kif18A motor domain has been solved in 2010 and it displays the typical kinesin fold described in paragraph 2.5.1. (Peters et al., 2010; Fig.2.7.A). In human cultured cells Kif18A accumulates specifically at the plus tips of k-fibres before anaphase onset, where it regulates microtubule dynamics through mild depolymerising activity and/or suppression of polymerisation dynamics (Du et al.,

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31 2010; Mayr et al., 2007; Stumpff et al., 2011; Varga et al., 2006). This activity is crucial for the suppression of chromosome oscillations and the establishment of a tight metaphase plate, as in the absence of Kif18A chromosomes undergo movements with higher amplitudes and fail to arrange in the middle of the cell, spindles are deformed and elongated and cells do not progress to anaphase due to activation of the spindle assembly checkpoint (Mayr et al., 2007; Stumpff et al., 2008). The kinesin is highly processive in vitro, meaning that it can walk for several micrometres and rarely falls off the microtubule track before reaching its plus tip where the molecule can dwell for long periods of time. This high processivity depends on an additional microtubule binding site in the C-terminus of Kif18A/Kip3 that also increases the microtubule affinity and on-time of the kinesins in vitro and is required for the efficient enrichment of Kif18A at the plus tip of k-fibres during mitosis (Mayr et al., 2011; Stumpff et al., 2011; Su et al., 2011;

Weaver et al., 2011). The mechanism by which members of the kinesin-8 family induce microtubule depolymerisation or a suppression of microtubule dynamics at plus tips is still debated. One model proposes that yeast Kip3 molecules dwell at the microtubule plus tip until new molecules arrive and “bump” into the dwelling ones, which then fall off the plus tip and take away one tubulin dimer during the process. Since the kinesin is very processive and one can assume that almost every molecule that binds the microtubule lattice will reach its plus tip and bump of a dwelling molecule, longer microtubules attached to kinetochores should accumulate more Kip3 than shorter ones, resulting in a more efficient shortening of these longer microtubules, a balancing of microtubule length and hence the arrangement of chromosomes in the middle of the mitotic cell (Varga et al., 2009). Whether this mechanism holds true also for Kif18A is still not completely clear (Fig.2.7.C). In vivo studies with chimeric constructs containing parts from different kinesins suggest that one important molecular

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32 element in Kif18A required to suppress chromosome oscillations is loop 2 in the motor domain.

Loop 2 is elongated in Kif18A compared to conventional kinesin and penetrates into the cavity above the interface of two tubulin heterodimers when the Kif18A motor domain is bound to microtubules (Kim et al., 2014; Peters et al., 2010; Fig.2.7.B). The involvement of loop 2 in the regulation of microtubule stability is appealing since the strong microtubule depolymerase MCAK (kinesin-13 member Kif2c, a kinesin with an internal motor domain) also depends on an elongated loop 2 to depolymerise microtubules (Asenjo et al., 2013; Moores et al., 2002; Wang et al., 2015b). On the other hand, loop 2 in Kif18A lacks the “KVD” motif that is essential for MCAK to depolymerise microtubules. An alternative function for loop 2, apart from induction of microtubule depolymerisation in Kif18A, might be that it provides a tighter association of the motor domain through interaction of its positively charged residues with the negatively charged microtubule lattice, thus enhancing processivity and plus tip accumulation (Kim et al., 2014;

Peters et al., 2010). The exact mechanism of how Kif18A depolymerises microtubules or supresses their dynamics in vivo remains a thrilling open question for future research.

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33 Figure 2.7. The kinesin-8 Kif18A. (A) Crystal structure of the human Kif18A motor domain from Peters et al., 2010 (PDB ID: 3LRE). The structure is rainbow colour-coded from blue (N-terminus) to red (C-terminus) and visible secondary structures are labelled. (B) Cryo-electron microscopy (cryo-EM) structure of the human Kif18A motor domain bound to microtubules with the crystal structures of the motor domain and α-β-tubulin fitted into the cryo- EM structure (Peters et al., 2010). Note that loop 2 (L2) of Kif18A that was not seen in the crystal structure appears as an ordered element when bound to the terminal region of α-tubulin. (C) Illustration of the accumulation of Kif18A molecules at the plus tips of k-fibres depending on the microtubule length; and simplified illustration of a Kif18A dimer. Note that the shorter microtubule accumulated less Kif18A molecules than the longer one. (D) Model for the network regulating Kif18A activity in early mitosis until metaphase from Hafner et al., 2014: Global and high Cdk1 activity results in inhibitory phosphorylation of Kif18A and its inability to accumulate at k-fibre plus tips in early mitosis. After proper bipolar attachment and stretching of kinetochores, the outer kinetochore gets separated from Aurora-B activity, allowing the binding of Protein phosphatase 1 (PP1) to the kinetochore. PP1 can dephosphorylate Kif18A allowing the kinesin´s efficient accumulation at k-fibre plus tips and the suppression of chromosome oscillations prior to anaphase onset.

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34 2.7.1. Kif18A is a suppressor of chromosome oscillations in metaphase

Oscillations of congressed chromosomes are thought to be required during prometaphase in order to detangle them from each other prior to segregation, because entangled chromatin fibres would lead to chromosomes that lag behind when pulled to opposite spindle poles, resulting in segregation errors during karyokinesis (Ke et al., 2009). However these oscillatory movements have to be suppressed shortly before anaphase onset to ensure the establishment of a “tight” metaphase plate. Like detangling, positioning of chromosomes in metaphase at the cell centre is a prerequisite for error-free chromosome segregation, because it ensures that all chromosomes have to roughly travel the same distance prior to cytokinesis. If some chromosomes would have to travel much longer than others, the spindle midzone would not be evenly established due to unequal distribution of the Ran-dependent, nucleating chromosome masses. Additionally, the likelihood of lagging chromosomes that could be entrapped by the contractile ring during cytokinesis would increase and eventually lead to daughter cell re-fusion due to the inability of the dividing cell to sever the midbody completely. Therefore the suppression of chromosome oscillations has to be precisely timed during late prometa- and metaphase. Recently, a regulatory network around Kif18A has been described that ensures accurate timing of the kinesin´s activity, resulting in the right timing of chromosome oscillation suppression (Hafner et al., 2014): Global and high Cdk1 activity in early mitosis leads to phosphorylation of Kif18A, which disables the kinesin to efficiently localise to the plus tips of k- fibres. When amphitelic end-on attachment of chromosomes is established, the phosphatase PP1 is recruited to kinetochores where it can dephosphorylate Kif18A, increasing its ability to localise at k-fibre plus tips and hence leading to the efficient suppression of chromosome oscillations in metaphase (Fig.2.7.D). Interfering with the ability of either Cdk1 or PP1 to

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35 modulate Kif18A activity ultimately results in more frequent chromosome segregation errors during anaphase, emphasising the importance of this regulatory network (Hafner et al., 2014).

Studies in living organisms suggest an important role of Kif18A in the germ line. Male Kif18A knock-out mice are infertile due to problems in the second meiotic division (Liu et al., 2010), which is in line with findings in Drosophila (Gatt et al., 2005; Savoian et al., 2004; Savoian and Glover, 2010). Additionally, pre-meiotic divisions in the mouse germ line depend on Kif18A.

Here, expression of a non-functional, motor-dead Kif18A version leads to an increase of cells arrested in mitosis with elongated spindles and unaligned chromosomes (Czechanski et al., 2015). Taken together, these results suggest that in multicellular organisms Kif18A activity is not required during all types of cell divisions (Liu et al., 2010), but is essential in tissues that depend on additional “fidelity” to ensure correct chromosome segregation, like for example the germ line. Along these lines, the high dependence of aneuploid cancerous human cell lines on Kif18A might reflect their need for higher fidelity of chromosome segregation during mitosis - a hypothesis that is further strengthened by the correlation of Kif18A overexpression in colorectal, breast and lung cancer and high rates of tissue invasion, metastasis and dissemination of these cancers in patients (Nagahara et al., 2011; Zhang et al., 2010).

2.8. Xenopus laevis - a model system to study spindle assembly and embryonic development The African clawed frog Xenopus laevis is a vertebrate model organism well established for studies on biochemical mechanisms of meiotic entry and resumption (Masui and Markert, 1971;

Sadler and Maller, 1985), processes required to initiate fertilisation (Rauh et al., 2005; Schmidt et al., 2005), embryonic development and cell cycle regulation (Tischer et al., 2012), and mechanisms of spindle assembly during M-phase. One major advantage of the Xenopus system

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36 is the availability of highly synchronised, large eggs (1 mm diameter) in big quantity. When injected with the hormone progesterone, one female frog can lay hundreds of eggs arrested at the metaphase of meiosis II, of which the isolated and nearly undiluted cytoplasmic fraction can be used for biochemical studies. Likewise, the ovulated eggs can also be fertilised in vitro to give rise to newly forming organisms and hence to study their embryonic development.

The open and therefore highly amenable cytoplasmic egg extract system can be utilised to study spindle assembly. Here, the M-phasic extract can be supplemented with demembranated sperm nuclei that contain a set of chromosomes and one centrosome, leading to the establishment of centrosome- and chromosome-containing spindles that can be visualised using fluorescence microscopy. Because it is an open system, the contribution of pathways/proteins to biological processes like spindle assembly can be assessed by immunodepletion and addition of recombinant proteins or respective coding mRNAs. Furthermore, completely new pathways can be discovered by combining the extract system with partial fractionation and mass spectrometry-based approaches. Studying spindle assembly in such an open system is a powerful tool that led to the discovery of several key regulators of this process, as for example the Ran-dependent microtubule nucleation around chromosome arms by the release of assembly factors like XMAP215 from inhibitory binding of importin β (Carazo-Salas et al., 1999;

Gruss et al., 2001), the identification of the kinesin Eg5 (Sawin et al., 1992), or the microtubule depolymerising kinesin-13 MCAK (Walczak et al., 1996).

During their early development, Xenopus embryos do not grow in size and undergo rapid cycles of cell divisions that lack gap phases and are only composed of DNA replication in S-phase and subsequent chromosome segregation and cell division in M-phase. As a consequence cell size within the early embryo is reduced drastically over the course of the first 11-12 divisions. Also,

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37 transcription is shut down in the early embryo and new proteins can only be synthesised from maternal mRNA templates that, along with energy sources like yolk proteins, were supplied by the mother. Since they are large in size and divide in a highly synchronous manner, in vitro fertilised Xenopus eggs and the resulting embryo are an ideal system to study the underlying biochemistry of cell cycle-related processes. The embryos are also highly resistant to external forces, enabling manipulation by microinjection of biochemically relevant amounts of protein, mRNA, or mRNA-targeting, stable nucleotide antisense probes like morpholinos.

2.9. Aims of this work

2.9.1. Generation of inhibitor-sensitive kinesin chimeras by transfer of an allosteric binding site

Kinesin motor proteins are often involved in rapid cellular processes. In order to study such processes, fast acting and reversible small molecule inhibitors represent ideal tools to ensure precise timing of inhibition. Since the number of available kinesin inhibitors is limited, alternative approaches to target these motor proteins in a highly spatio-temporal manner could help to mitigate the lack of tailored and potent small molecule inhibitors. Therefore the aim in the first part of this work was to establish a transferrable inhibition system for kinesins. Loop 5 in the kinesin Eg5 is the binding region of several well characterised allosteric inhibitors and structural information about some of the enzyme-inhibitor complexes are available. Also, secondary and tertiary structure among kinesin motor domains is highly conserved, allowing the generation of functional motor domain chimeras. It was therefore decided to transfer loop 5 as an allosteric binding site from Eg5 into other kinesins and test the generated kinesin chimeras

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38 for their functionality, as well as their sensitivity towards one loop 5-directed compound, S-trityl cysteine (STLC) in vitro by employing an enzyme coupled assay system.

2.9.2. Biochemical and functional characterisation of Xenopus laevis Kif18A

It is established in a variety of model systems that members of the kinesin-8 family are required for proper chromosome segregation by regulating microtubule stability and hence spindle morphology throughout different stages of M-phase. Much is known about the importance of kinesin-8 members in yeast and transformed cultured cell systems during mitosis, but there is also rising evidence for a function in the reductional meiotic divisions in higher eukaryotes.

Recently it was proposed that the family member Kif18A is a “fidelity factor” for chromosome segregation during mitotic divisions in the mouse germ line. Both, errors during divisions in the germ line and during embryonic development are detrimental to the fate of newly forming organisms. This raised the question whether zygotic cell divisions, which follow meiosis and fertilisation of the egg, also depend on the activity of Kif18A. In order to address this question, Kif18A had to be identified in the African claw frog Xenopus laevis, an ideal model organism to study embryonic development. Subsequently, the role of the kinesin was to be assessed by combining the strength of biochemical in vitro analysis with functional assays in the egg extract system and developing embryos.

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39

3 Results

3.1. Transfer of an allosteric binding site renders Kif3A and Kif4A sensitive towards STLC, a small molecule inhibitor of Eg5

One aim of this thesis was to generate kinesin mutants that are sensitive towards an allosteric Eg5 inhibitor. This part of the work describes the successful generation of such chimeric enzymes in which stretches of the motor domain have been replaced with parts from Eg5 in order to confer sensitivity towards the Eg5 inhibitor S-trityl cysteine (STLC). Two helices and a loop in Eg5´s motor domain were sufficient to confer sensitivity towards STLC.

3.1.1. The transfer approach of Eg5´s loop 5 into other kinesins

The Drosophila homologue of Eg5, Klp61F, is not inhibited by STLC. It has however been reported that the exchange of critical residues within the STLC binding site in Klp61F resulted in its inhibition by the compound (Liu et al., 2011). Intrigued by these findings, it was decided to transfer the STLC binding site from Eg5 into other kinesin motor domains in order to confer sensitivity towards the compound (Fig. 3.1.1.A). In detail, both helix 2 together with loop 5 and helix 3 up to the switch I region of Eg5 were transferred into motor domains of Kif3A and Kif4A (Fig.3.1.1.C). As these three structural elements reconstitute the whole binding bed of STLC (henceforth conferred to as the STLC binding cluster), it was assumed that if correctly positioned, this newly formed binding site should interact with STLC and also lead to an inhibition of the kinesins. This theory was further encouraged by the similar fold among kinesin motor domains, as illustrated by the overlay of human Eg5 and murine Kif4A heads (Fig.3.1.1.B).

Both helix 2 and 3 have the same length in the trinucleotide-bound state and show similar steric

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40 arrangements in Kif4A and Eg5. The biggest divergence can be found in loop 5, which is longer in Eg5.

Figure 3.1.1. The transfer approach of Eg5´s loop 5 into other kinesins. (A) Illustration of the allosteric binding site transfer approach. The known and well characterised binding site of a small molecule inhibitor (yellow) in a kinesin (red) is transferred into another kinesin (blue) for which no small molecule inhibitor is available. The resulting chimeric kinesin (blue and red) harbours the inhibitor binding site and is therefore susceptible to the compound. (B) Overlay of human Eg5 and murine Kif4A motor domains in a trinucleotide (light red and green)-bound state with focus on helices 2, 3 and loop 5 (binding site of STLC in Eg5 in red, corresponding region in Kif4A in light blue). Note that the helical elements are arranged in almost exactly the same angle and position in both structures. Structures from Parke et al., 2010 (Eg5) and Chang et al., 2013 (Kif4A). (C) Alignment of wildtype (wt) and chimeric (C) motor domains from Homo sapiens (hs) Eg5, Kif3A and Kif4A using T-Coffee (Notredame et al., 2000). Secondary structure elements extracted from the Eg5 crystal structure are marked on top of the alignment line. Bold letters represent residues important for STLC binding in Eg5 (Liu et al., 2011). Colour coding: sequence stretches in Eg5 that are highlighted in the structure from (B) are in red. Stretches in Kif3A (hsKif3A wt: green) and Kif4A (hsKif4A wt: blue) that were replaced with Eg5 sequences in the chimeras (hsKif3A C and hsKif4A C) are also in red.

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41 3.1.2. Biochemical characterisation of the chimeric kinesin motor domains

The in silico-designed chimeric motor domains of Kif3A and Kif4A that contain Eg5´s STLC binding cluster (chimeras, abbreviated C), as well as wildtype motor domains (abbreviated wt) of Eg5, Kif3A and Kif4A, were obtained using Gibson cloning. The constructs were expressed in E.coli and enzymes were recovered from the soluble lysate using Nickel affinity chromatography (Fig.

3.1.2.B). To directly test the activity of the kinesins, an enzyme-coupled assay system was employed. ATP turnover can be measured using the activity of two additional enzymes, pyruvate kinase and lactate dehydrogenase, respectively, leading to the oxidation of NADH to NAD⁺ during ATP consumption. NADH absorbs at a wavelength of 340 nm, whereas NAD⁺ does not, resulting in a reduction in the absorption at 340 nm when ATP is consumed (Fig. 3.1.2.A).

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42 Figure 3.1.2. Biochemical characterisation of the chimeric kinesin motor domains. (A) Illustration explaining the principle of the applied enzyme-coupled assay. ATP is hydrolysed by the kinesin. ADP is recycled to ATP by pyruvate kinase-mediated conversion of phosphoenolpyruvate to pyruvate. Pyruvate is subsequently reduced to lactate by lactate hydrogenase and the oxidation of NADH to NAD⁺. NADH absorbs at 340 nm, NAD⁺ does not. A decrease of A340 therefore is a direct measurement for ATP turnover in the system. (B) Coomassie brilliant blue (CBB) staining of a SDS-polyacrylamide gel showing the five different purified motor domains hsEg5 wildtype (wt), hsKif3A and hsKif4A wildtype (wt) and chimeras (C), respectively. (C) Relative molar activity of the respective motor domains under fully activating ATP and microtubule (MT) concentrations in the enzyme-coupled assay. Wildtype activity was set to 100% for n=3 different measurements per condition. (D) Summary of Km (ATP) and K1/2 (MT-microtubules) values obtained from the diagrams E-J using Michaelis-Menten or sigmoidal (only for (I), Kif3A C) fit in GraphPad. (E- F) ATP titration for kinesins from (B), using constant microtubule (saturated) and kinesin concentrations; (H-J) Titration of microtubules (MT) for kinesins from (B), using constant ATP (saturated) and kinesin concentrations in the enzyme-coupled assay. Error bars represent standard deviation from three experiments. Fit as described in (D).

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43 In order to estimate the overall activity of chimeric motor domains, their molar activity in respect to the wildtype enzymes was determined. Both Kif3A and Kif4A chimeras displayed a drop in activity of approximately 20% compared to the respective wildtype under saturating substrate (ATP) and cofactor (microtubules) conditions (Fig. 3.1.2.C). This was appreciated as a positive result after the significant sequence exchange in these proteins. ATP concentrations at half maximal enzyme activity (Km) were in the lower micromolar range for all five purified motor domains and could be fitted assuming Michaelis-Menten kinetics, implying functionality of all five enzymes (Eg5wt, Kif3A wildtype and chimera, Kif4A wildtype and chimera; Fig. 3.1.2.D and E-G). Particularly, the Km values of wildtype and chimeric enzymes were nearly equal for both Kif3A and Kif4A (Fig. 3.1.2.F and G). All kinesin motor domains displayed microtubule (MT)- stimulated ATPase activity, as indicated by K1/2 values in the lower nanomolar range, which followed hyperbolic stimulation kinetics (Fig. 3.1.2.D and H-J). Hence, the important regulation of kinesin activity by its cofactor and track, the microtubule, is still functional in the generated mutants. The Kif4A chimera displayed very similar stimulation kinetics by microtubules compared to the wildtype (22 vs. 21 nM; Fig. 3.1.2.D and J). However, exchanging Kif3A´s helix 2, loop 5 and helix 3 in order to introduce the STLC binding cluster led to a significant reduction in microtubule-stimulated activity. The K1/2 value was approximately six times higher than in the wildtype (4 vs. 23 nM) and also followed a more sigmoidal stimulation (Fig. 3.1.2.D and I). This indicates a lower affinity towards microtubules of the Kif3A chimera compared to the wildtype, which might be manifested in a higher Koff rate, a lower Kon rate or a combination of both.

Functional Characterisation of Kif18A in Xenopus laevis and Generation of Inhibitor-Sensitive Kinesin Chimeras