The microtubule cytoskeleton
The rapid reorganization of cellular content in cycling cells is largely dependent on a system of protein fibers, called the cytoskeleton. Just as a body skeleton it serves as a scaffold to a cell, organizing its spatial architecture and giving it physical resistance. However, the cytoskeleton is a very dynamic structure composed of filaments that keep modifying their length and are able to quickly reorganize into different configurations.
The cytoskeleton is composed of three types of proteinaceus filaments: actin microfilaments, intermediate filaments and microtubules. Microtubules are the thickest of cytoskeleton components. Their lattices are polymers of α-‐ andβ-‐tubulin, 25nm in diameter that can grow even up to hundreds of micrometers. They arrange into arrays to form various microtubule-‐based structures and govern intracellular transport. They are organized from Microtubule Organizing Centers (MTOCs), centrosomes being the most frequent example. I will first discuss the structure and dynamics of the microtubule cytoskeleton and then move to its organization throughout the cell cycle.
Microtubule lattices are dynamic polymers and their subunits are added and removed in interconverting processes called polymerization and depolymerization. Their building blocks are heterodimers of α-‐ andβ-‐tubulin (Fig.4). The two isoforms of tubulin are homologous but not identical and in a mixture dimerize immediately and never come apart. Both contain a site where they bind GTP. They are called N-‐site (for Non-‐exchangeable) in α-‐tubulin and E-‐site (for Exchangeable) in β-‐tubulin since only β-‐tubulin hydrolyzes GTP during polymerization and exchanges GDP to GTP during depolymerization. GTP on the α-‐tubulin subunit remains enclosed in the microtubule lattice as it polymerizes. The energy from GTP hydrolysis on β-‐tubulin allows for non-‐ equilibrium polymerization dynamics (see below).Heterodimers assemble with each other in a head-‐to-‐tail fashion and form long α-‐β-‐tubulin protofilaments (Fig. . 4)
Monomers of α-‐tubulin can also interact with α-‐tubulin monomers from another protofilament and the same is true for β-‐tubulin. Due to such lateral interactions between adjacent protofilaments, they assemble together andform a sheet-‐like structure that elongates and subsequently encloses in a tubular microtubule. In vivo assembled microtubules are composed of 13 laterally eractingint protofilaments (Tilney et al., 1973). However, from 10 until 15 protofilaments can make up a microtubule in in vitro assays(Pierson et al., 1978) suggesting that it is not the structural requirement of tubulin subunits that imposes the 13-‐fold symmetry, but other factors must contribute to microtubule assembly in vivo. Notably, microtubules assembled in vitro from a centrosome are all composed of 13 protofilaments (Evans et al., 1985), strongly suggesting that the nucleating structure controls the number of protofilaments within a microtubule.
From the head-‐to-‐tail assembly of heterodimers stems an essential feature of microtubules, which is their polarity. It is manifested by the fact, that microtubules have a faster growing end, called the plus-‐end, that has subunits of β-‐tubulin exposed at the tip, and a slower growing end, called the minus-‐end, with α-‐tubulins at the tip. The polymerization of a microtubule lattice is a two-‐step process: first a GTP-‐bound subunit is added to the end of a microtubule, and then GTP is hydrolyzed and a phosphate (Pi) is released during polymerization or just after (Fig. 4 and. The5) disassembly comprises only detachment of a GDP-‐bound dimer with a fast rate. Curved protofilaments are observed at the end of a depolymerizing microtubule (Fig. 5). What seems to be determining the dynamicity of microtubule polymers is the concentration of free tubulin dimers that is available. Above a critical concentration (Cc) of tubulin, dimers are assembled on a microtubule, while below this Cc concentration, they are dissociated. Detailed study of nucleotide-‐bound tubulin association and dissociation rates as a function of tubulin concentration aim to understand how microtubules maintain prolonged phases of polymerization or depolymerization and why these phases interconvert (Walker et al., 1988; Erickson and O’Brien, 1992). In vivo, the newly nucleated microtubules are anchored to an MTOC via their minus ends. Polarity is central to motor protein activity, as they are usually able to move along the microtubule lattice in one unique direction using the energy from ATP hydrolysis. In general, motor proteins from the kinesin family (Vale and Fletterick, 1997) move towards the plus end, and dyneins (Schroer et al., 1989) towards the minus end.
Due to their hollow, rod structure, microtubules have an intrinsic resistance to bending and squashing. However, the non-‐covalent nature of polymer joints provides them with the ability to quickly polymerize and depolymerize to change their length. In fact, both tips of a microtubule lattice are constantly interconverting between two types of events: catastrophe, which is an abrupttransition from polymerization to
depolymerization, and rescue, which is the reverse process (Fig.5). This non-‐ equilibrium dynamics is called “dynamic instability” (Mitchison and Kirschner, 1984) and although it is energetically costly, it has been evolutionarily conserved implying its importance in the functionality of the microtubule cytoskeleton. First, it enables its rapid reorganization: it is much easier for small independent units to diffuse between cell compartments and reconstruct microtubule arrays in response to internal and external factors. Second, it is a undationfo for microtubule-‐dependent force generation, as polymerization of microtubules can result in pushing of objects, and depolymerization combined to a motor protein activity is able to pull on objects. Dynamic instability is the resultant of the rates of polymerization and depolymerization, and the frequency of catastrophe and rescue events. Since 1984, when Mitchison and Kirschner first described the mechanism and postulated this term, it has been confirmed both in vitro andin vivo and was recognizedas the main mechanism governing thedynamics of microtubules. Supplementary concept for describing the dynamic behavior of the polymer, called treadmilling, comes from the observation that microtubules at steady state continuously incorporate new tubulin dimers (Margolis and Wilson, 1978). When conditions allow for it, at one end of treadmilling polymer tubulin subunits are constantly assembled, while at the opposite end they are constantly lost at a balanced rate, which results in a forward movement of the polymer without any change in its length.
Microtubule associated proteins
An increasing number of proteins has been identified thatinteract with microtubules. They are globally referred to as Microtubule Associated Proteins (MAPs), but subgroups of very distinct functions can be distinguished. In eukaryotes, MAPs include microtubule-‐based motor proteins, structural MAPs, proteins that associate with the plus tip of microtubules, centrosome associated proteins and MAPs that are enzymatically active. Many of them regulate microtubule dynamics by stabilizing the protofilaments, or conversely promoting the catastrophy or the severing of microtubule lattices. Some MAPs have repetitive domains, which allow them to interact with many microtubules at the same time and locally induce common regulation and interdependent organization. Below, I will highlight the activity of a few groups of MAPs that are relevant in the context of my PhD.
Within cells, microtubule growth is highly accelerated compared to in vitro experimental conditions with purified tubulin and GTP solutions. This is achieved through activity of microtubule stabilizing MAPs; Fig. 6 (Cassimeris, 1993; Desai and Mitchison, 1997).
Plus-end tracking proteins (+TIPs)
A group of proteins that associates with the faster growing end of microtubules are the +TIPs. They are structurally and functionally divergent, but they all bind to the plus-‐end directly or through a partner. Once recruited, most +TIPs stabilizethe microtubule lattice and favor polymerization by decreasing catastrophe frequency. The first identified +TIP protein was the cytoplasmic linker protein, CLIP-‐170 (Perez et al., 1999). Since its discovery, more than 20 families of +TIP proteins have been described (Yu et al., 2011; Larsen et al., 2013).
The end-‐binding (EB) family of proteins is capable of direct interaction wit microtubules (Hayashi and Ikura, 2003). They also provide a platform for association of other +TIPs that don’t contain a microtubule-‐binding domain themselves, like CLIP-‐170 (Perez et al., 1999; Goodson et al., 2003). CLASPs are proteins that promote addition of tubulin dimers at the plus end and inhibit catastrophe. During mitosis, they are among proteins involved in microtubule stabilization at the kinetochores (Fig. 6); (Cheeseman et al., 2005; Maiato et al., 2005; Pereira et al., 2006).
HURP (Hepatoma Upregulated Protein) is a MAP that stabilizes kinetochore-‐ microtubule attachments in mitosis (Koffa et al., 2006; Silljé et al., 2006)and is essential for meiotic spindle formation in mouse oocytes (Breuer et al., 2010). It is a Ran effector (see 126.96.36.199.2.) required for assembly of a central microtubule domain in the vicinity of chromosomes, where it accumulates via kinesin-‐5 activity. In mouse oocytes, this domain acts as a scaffold for poleward sorting of MTOCs and chromosome alignment within the metaphase plate and is essential for spindle establishment and maintenance.
Several kinesin families (microtubule motors, see below) are known for their microtubule destabilizing activity. They can promote dissociation of tubulin dimers by increasing the frequency of catastrophe and ingfavorthe curved protofilament conformation (Fig. 5).The founding member of kinesin-‐13 family, MCAK is able to remove GTP-‐bound tubulin dimers using energy from ATP hydrolysis (Fig. 6); (Hunter et al., 2003). It accelerates microtubule depolymerization 100-‐fold and induces the curved protofilament conformation. During mitosis it regulates the length of the spindle (Walczak et al., 1996; Goshima and Vale, 2003; Domnitz et al., 2012) and at anaphase promotes dissociation of tubulin dimers at the kinetochore, as kinetochore-‐linked microtubules shorten to separate sister chromatids (Maney et al., 1998; Rogers et al., 2004).
Other examples of destabilizing MAPs are katanin, spastin and fidgetin. They are microtubule severing enzymes that function in microtubule release from the centrosome, disassembly of microtubule interphase network at mitosis onset, maintaining spindle structure and chromosome alignment in metaphase and chromatid separation at anaphase (Hartman et al., 1998; Ahmad et al., 1999; Zhang et al., 2007; Roll-‐Mecak and McNally, 2010; Loughlin et al., 2011; McNally et al., 2014).
Microtubule-‐based motor proteins are MAPs able to move along the lattice of microtubules. They use energy from ATP hydrolysis to generate the force necessary for processive movement and are able to bind and transport a variety of cargos. They comprise two large families of enzymes, dyneins and kinesins(Holzbaur and Vallee, 1994; Vale and Fletterick, 1997).
Dyneins are minus-‐end directed microtubule motors (Fig. 7).They form functional complexes around force-‐generating subunits, called heavy chains (DHC).Dyneins belong to the AAA+ superfamily (ATPases Associated with diverse Activities); (Neuwald et al., 1999), as DHC is composed of six AAA+ modules that fold into a ring (Fig. 7; blue donut-‐shape). DHC also contains domains responsible for binding both to a microtubule and to a cargo. They further assemble together with three classes of associated subunits: intermediate chains (DIC), light intermediate chains (DLIC) and light chains (DLC).For full activity, dynein requires another multisubunit protein complex, dynactin, which binds to the DIC. Dynactin acts as an “adaptor” that expands the range of cargos that dynein can move and increases processivity of dynein; for review see Kardon and Vale (2009).
During interphase, cytoplasmic dynein mediates the transport of vesicles, little organelles and mRNA. As a minus-‐end directed motor, it moves processively towards the center of the cell where microtubules are anchored at MTOCs. Dynein localizes at the cortex where it exerts pulling forces on microtubules and can thus participate in microtubule-‐based organelle positioning, including the nucleus(Tanenbaum et al., 2011).
Table of contents :
1. The cell cycle
2. The microtubule cytoskeleton
2.1. Microtubule structure
2.2. Microtubule dynamics
2.3. Microtubule Associated Proteins
2.3.1. Stabilizing MAPs
188.8.131.52. Plus-‐end trancking proteins (+TIPs)
2.3.2. Destabilizing MAPs
2.3.3. Motor proteins
2.4. Microtubules during the cell cycle
2.4.1. Microtubules in interphase
2.4.2. Microtubules in mitosis
184.108.40.206. Pathways of microtubule spindle assembly
220.127.116.11.1. Centrosome pathway
18.104.22.168.2. Chromosome pathway
22.214.171.124.3. Augmin pathway
2.4.3. Microtubules in anaphase
2.4.4. Microtubules in cytokinesis
2.5. Centers of microtubule organization
2.5.1. The centrosome
2.5.2. Non-‐centralized microtubule organization
3. Regulation of centrosome-‐linked activities
3.1. Centrosome cycle
3.2. PCM formation
3.3. Regulation of centrosome cycle
3.3.1. License to duplicate and cell cycle control of centriole duplication
3.4. Centrosome cycle of genome stability
3.5. PLK4 as a master regulator of centriole duplication
3.5.1. PLK4 structure
3.5.2. PLK4 regulation
3.4.3. PLK4 localization to the centrosome and function in centriole assembly
3.4.4. PLK4 role in centriole over-‐duplication and de novo formation
3.4.5. PLK4 activity in acentriolar cells
4. Mammalian female meiosis in the absence of centrioles
4.1. Disapearance of centrioles
4.2. Protracted prophase I arrest
4.2.1. Control of prophase I arrest by cAMP
4.2.2. Regulation of Cyclin B levels by APC/C
4.2.3 Governing the microtubules in prophase I in the absence o centriole
4.3. Meiotic maturation
4.3.2. Acentriolar assembly of the first meiotic spindle
4.3.3. Spindle bipolarization and MTOC sorting to the poles
4.3.4. Chromosome congression and k-‐fibers assembly
4.3.5. The SAC in female meiosis
4.3.6. Spindle migration to the cortex and chromosome segregation
4.3.7. Polar body extrusion and metaphase
1. Rebuilding MTOCs upon centriole loss during mouse oogenesis
2. Plk4 regulation of acentriolar MTOC assembly is critical for meiosis in the mouse oocyte
Towards understanding of PCM assembly Regulation of PCM recruitment to acentriolar MTOCs
Centriole elimination in the mouse oocyte