Microtubules Regulate Cell Migration and Neuronal Pathfinding
While many cell types are able to generate cellular movement through the action of the actomyosin cytoskeleton alone, microtubules are important for establishing and maintaining polarity, regulating the force-generating machinery and cell adhesion. Therefore, directionally persistent cell migration and neuronal pathfinding often require microtubules.
The microtubule cytoskeleton itself is organised asymmetrically to allow differential regulation of the migration machinery at the front and the rear of the cell. Microtubules position organelles such as the nucleus, the centrosome and the Golgi. Transport of mRNAs, vesicles, receptors and signalling components to the cell edges occurs along microtubules. These cargoes in turn support force generation by the actin cytoskeleton, act as a source of membrane lipids and regulate polarity signalling, adhesion, cell-cell communication and chemical gradient sensing. Microtubules themselves and especially the dynamic plus ends act as signalling platforms to control adhesion turnover and membrane protrusion. The rapid turnover of microtubules allows cells to quickly adapt to extracellular signals and change migration direction in response to guidance cues. Microtubule dynamics and organisation are in turn controlled by cortical cues. These feedback mechanisms ensure robustness and adaptation to environmental influences.
Given the fundamental importance of cell migration for embryonic development, the immune system and wound healing, impaired microtubule function leads to birth defects and diseases. Likewise, drugs targeting microtubules are routinely used to prevent excessive cell migration in cancer metastasis and chronic inflammatory diseases.
Diseases directly associated with cell migration and microtubule function
Microtubule structure and stability:
Mehlen and Puisieux (2006)
Immune cell migration
Chia et al. (2008)
Infiltration of immune cells into the brain; multiple sclerosis-like phenotype
Experimental autoimmune encephalomyelitis
O’Sullivan et al. (2013)
Neuronal migration defects
Malformation of cortical development
Neuronal migration defects
Poirier et al. (2007)
Neuronal migration defects
Jaglin et al. (2009)
Neuronal migration defects
Breuss et al. (2012)
Neuronal migration defects
Keays et al. (2007)
Neuronal migration defects
Malformation of cortical development
Poirier et al. (2013)
Neuronal migration defects
Neuronal migration defects
Impaired neuronal network formation
Blood vessel formation
Tumour angiogenesis, metastasis
Increased vessel density in tumours
Sun et al. (2013)
Microtubule length and array control:
Increased microtubule severing
Sapir et al. (2012)
Sperm motility defective
Male fertility defect
O’Donnell et al. (2012)
Impaired microtubule severing
Draberova et al. (2011)
Enhanced cell motility and invasiveness
Motor proteins and their regulation:
Movement of organelles, retrograde trafficking affected
Charcot-Marie-Tooth disease type 2; several neurological symptoms
Willemsen et al. (2012)
Impaired dynein function
Lissencephaly, Miller-Dieker syndrome
Neuronal migration defects
Malformation of cortical development
Poirier et al. (2013)
Centrosomal function impaired
Schizophrenia, depression, bipolar disorder
Centrosomal satellites defective
Kamiya et al. (2008)
Centrosomal satellites defective
Defective cilia; defects in migration cause craniofacial dysmorphia
Tobin et al. (2008)
Excess centrosomes in interphase
Angiogenesis; defective vessel sprouting
Kushner et al. (2014)
Cell polarity signalling:
Stability of microtubules in the uropod of neutrophils
Kumar et al. (2012)
Microtubule-regulated adhesion turnover defective
Delayed skin healing
Wu et al. (2008)
Cell adhesion by cadherins affected
Tumour development/metastasis in colorectal cancer
Faux et al. (2004)
Cells can migrate in various different modes that depend on the environment they are in and on the cell type. On a flat surface such as a plastic dish in culture or the surface of a muscle fibre or endothelial sheets in vivo, cells move in a mesenchymal mode with adhesion to the surface being a crucial aspect of migration. Moving through dense 3D matrices or other confined spaces requires only little adhesion as under these conditions contractile forces that drive amoeboid or blebbing motion can generate forces and traction at the same time. Cells can migrate as individual cells or as collectives, and they can also switch between different types of migration (Friedl and Gilmour 2009). Such a change occurs, for example, during epithelial-mesenchymal transition, a process where cancer cells undergo dedifferentiation from a tissue collective to a more single-cell-like behaviour and acquire the ability to metastasise (Friedl and Wolf 2003).
Protrusion. This involves the cell membrane to be pushed forward by cytoskeletal polymerisation.
Adhesion. Forces that the cytoskeleton generates must be transmitted to the underlying substratum while regulating the turnover (lifetime) according to the spatial cues (strong attachment at the front, weakening attachment at the rear).
Contraction. Actin and myosin generate contractile forces to move the cell body forward.
Retraction. Substrate adhesion at the rear must be released and the rear end of the cell brought forward.
This classic model describes the series of events needed to propel a cell forward. The importance of each of these aspects differs depending on the type of migration, e.g. mesenchymal migration strongly depends on attachment, while amoeboid migration does not (for details, see Lammermann and Sixt 2009). In order to achieve persistent directional motility of a cell, there are essential requirements that need to be met: First, cell polarity needs to be established. Next, the cytoskeleton needs to be arranged so that forces are generated in the different parts of the cell that allow protrusion at the front and retraction at the rear. These forces need to be transferred to the underlying substrate with the help of adhesive contacts, either to the extracellular matrix (ECM) or to neighbouring cells. Additional tasks are added where cells migrate in clusters, as contacts and communication between the migrating cells need to be maintained at all times.
Although migration is often regarded as a purely actin-driven process, microtubules have fundamental roles in the regulation of different aspects of the complex task of moving a cell forward. However, the exact involvement of microtubules in migration is strongly dependent on the type of cell and its environment. Leaving aside protists, whose motility depends entirely on microtubules organised into cilia, it appears that in small cells, such as neutrophils (Dziezanowski et al. 1980; Niggli 2003), T cells (Takesono et al. 2010) or fish keratinocytes (Euteneuer and Schliwa 1984), microtubules are dispensable for efficient migration, even if some aspects of migration require microtubules (Stramer et al. 2010; Vogl et al. 2004, Fig. 6.1e). This was demonstrated in experiments using microtubule-depolymerising drugs, e.g. nocodazole or colcemid. When small cell types were treated with these drugs, their migration was hardly impaired or even stimulated (Euteneuer and Schliwa 1984; Niggli 2003). Yet when the experiment was repeated on larger cell types, such as fibroblasts, neurons, astrocytes or cancer cells, the effects on migration ranged from loss of directionality and cell polarity and reduction of speed to complete inhibition of cell locomotion (Etienne-Manneville 2004; Ganguly et al. 2012; Liao et al. 1995; Vasiliev et al. 1970; Xu et al. 2005). One idea is that diffusion or actin-based transport can efficiently compensate for loss of microtubules in small but not in larger cells (Kaverina and Straube 2011; Keren et al. 2008).
6.2 Microtubule Organisation in Migrating Cells
The Golgi complex is usually positioned close to the centrosome (Kupfer et al. 1983; Pouthas et al. 2008) and nucleates a large number of almost exclusively front-directed microtubules from the trans-Golgi network (Chabin-Brion et al. 2001; Efimov et al. 2007; Rivero et al. 2009; Fig. 6.3). As the centrosome and the associated microtubules organise the Golgi apparatus, this coupling of centrosomal positioning and Golgi-mediated nucleation of microtubules increases the front-biased orientation of microtubules in the cell (Vinogradova et al. 2012). An extreme example of higher microtubule density extending towards the front occurs in the very long, but narrow, lamellipodia of migrating granule cell neurons (Umeshima et al. 2007).
Only a few of all microtubules growing towards the leading edge of the cell actually reach the plasma membrane. These are so-called “pioneer” microtubules (Etienne-Manneville 2013). Most other front-oriented microtubules terminate near the actin-rich regions of the cortex, but do not touch the expanding membrane at the front. It is thought that retrograde flow from the actin filaments prevents these microtubules from reaching the membrane (Waterman-Storer and Salmon 1997). “Pioneer” microtubules withstand expulsion by actin retrograde flow by anchorage to the membrane (Etienne-Manneville et al. 2005). The observation that “pioneer” microtubules show extensive tubulin modifications supports the idea of increased longevity of this microtubule population (Bulinski and Gundersen 1991; Gundersen and Bulinski 1988). A similar arrangement is found in axons, where only a subset of microtubules enters the peripheral domain of the growth cones (Fig. 6.2c).
In differentiating neurons, the cell body will no longer move forward, but the growth cones at the tips of the extending neurites structurally and functionally resemble the lamella of migrating cells. Growth cones are able to continue to grow in the absence of microtubules, but the sensing of chemical gradients of guiding cues is impaired and directional growth is lost (Williamson et al. 1996). The directionality of growth is determined by highly localised actin protrusion and adhesiveness on one side of the growth cone against the other (Vitriol and Zheng 2012; Fig. 6.2c). This correlates with changes to the microtubule array: Microtubules are stabilised on the protruding and destabilised on the collapsing side, possibly through the action of APC (Buck and Zheng 2002; Zhou et al. 2004). One idea is therefore that microtubules direct the delivery of vesicles, mRNAs and GTPase activators to the growing side of the axon tip. The microtubule organisation in growth cones is dominated by front-directed microtubules that grow from the neurite into the growth cone (de Anda et al. 2005). Most of these microtubules do not extend all the way from the centrosome or Golgi network. Non-centrosomal microtubule nucleation occurs throughout the axon and dendrites (Stiess et al. 2010; Yau et al. 2014). In addition, severing enzymes such as Katanin or Spastin release microtubules from their anchoring at the centrosome, thereby enabling motor-driven transport of microtubules into neurites (Liu et al. 2010; Myers and Baas 2007; Yu et al. 2008). Advancing microtubules into the peripheral domain is then mediated by molecular motors of the kinesin-5 and kinesin-12 family (Nadar et al. 2008; Liu et al. 2010). In some migrating cells, the release of microtubules from the centrosome and cytoplasmic transport has also been observed, suggesting that similar mechanisms for microtubule reorganisation exist in migrating cells (Abal et al. 2002; Jolly et al. 2010).
In addition to the release of microtubules from their nucleation site, severing proteins also allow the destruction or amplification of microtubule subpopulations and can therefore modify the number of microtubules in a given orientation (Lacroix et al. 2010; Lindeboom et al. 2013; Sudo and Baas 2010). Cutting the microtubule lattice will produce two microtubules with the same orientation that either rapidly depolymerise or are stabilised and grow. Newly created minus ends are stabilised by CAMSAP family proteins. Depletion of CAMSAP2 results in a reduction in posttranslationally modified microtubules, cell polarity and directional cell migration (Jiang et al. 2014), suggesting that the stabilisation of non-centrosomal microtubules and the amplification of front-directed microtubules through collaboration of severing enzymes and minus end stabilisers are important for the asymmetric microtubule arrangement in motile cells.
Katanin localises to the leading edge of migrating human and Drosophila S2 cells and negatively regulates migration of these cells in vitro (Zhang et al. 2011). Katanin appears to be enriched at sites of filopodia formation (Liu et al. 2008), and increased amounts of Katanin subunits have been linked to more aggressive migratory behaviour in prostate cancer cells (Ye et al. 2012). Similarly, inhibition of Katanin subunits leads to migration impairment in mouse neurons and rat epithelial cells (Sudo and Maru 2008; Toyo-Oka et al. 2005). Uncontrolled function of Katanin and Spastin leads to aberrant numbers of microtubules in neurons, which has been linked to a number of diseases such as hereditary spastic paraplegia or Alzheimer’s disease (Errico et al. 2002; Sudo and Baas 2011), causing general defects in microtubule-mediated transport.
In addition to increased nucleation of microtubules towards the front of the cell and potential amplification mechanisms by severing enzymes, differences in microtubule stability contribute to the asymmetry of the microtubule cytoskeleton. Tubulin acquires posttranslational modifications in long-lived microtubules. In migrating cells, a front-directed accumulation of microtubules containing acetylated and detyrosinated tubulin is often observed (Gundersen and Bulinski 1988; Umeshima et al. 2007), suggesting that front-directed microtubules are stabilised, thus further exacerbating microtubule asymmetry.
It is thought that the asymmetry in the microtubule array allows preferential traffic of cargoes to the front of the cells (Fig. 6.3; Bachmann and Straube 2015). Important cargoes for cell migration are actin and Arp2/3 mRNA (Lawrence and Singer 1986; Mingle et al. 2005), post-Golgi carriers (Miller et al. 2009; Yadav et al. 2009) and recycling endosomes (Palamidessi et al. 2008). As posttranslational modifications of tubulin can serve as guidance cues for microtubule motor proteins, efficient front-directed transport can be achieved by a combination of increased number and selective stabilisation and modification of microtubules to the leading edge. Track selectivity has been demonstrated for kinesin-1s, kinesin-2s and dynein (Dixit et al. 2008; Sirajuddin et al. 2014) in vitro, and there is some evidence that this is also the case in cells (Cai et al. 2009; Ghosh-Roy et al. 2012; Huang and Banker 2012; Jacobson et al. 2006; Reed et al. 2006). In mature neurons, this property of kinesins to preferentially bind differentially modified tubulin is exploited to selectively target cargo specifically to axons or dendrites (Burack et al. 2000; Jenkins et al. 2012). Likewise, a preference for transport to and accumulation at the rear of migrating cells has been shown for the kinesin-3 Kif1C, which is negatively regulated by tubulin acetylation (Bhuwania et al. 2014; Theisen et al. 2012).
Posttranslational modifications of tubulin also regulate the activity of Katanin and Spastin and the binding affinities of microtubule-associated proteins (MAPs) such as Tau. While acetylation and polyglutamylation of tubulin increases severing activity, decoration of the microtubule lattice with Tau protects microtubules from severing (Lacroix et al. 2010; Sudo and Baas 2010). Abnormal regulation of Tau has been associated with disease progression, most notably with neurodegenerative diseases such as dementia (Lee and Leugers 2012). Thus complex feedback loops involving chemical modification and modification-sensitive MAPs modulate the asymmetric microtubule network in migrating cells.
6.3 Spatial Regulation of Microtubule Dynamics
As mentioned above, differences in microtubule dynamics at the front and rear of the cell contribute to the asymmetry in the microtubule organisation. Cells express an arsenal of microtubule regulators that tightly control the assembly and disassembly of microtubules (van der Vaart et al. 2009). In cells, microtubule catastrophe occurs almost exclusively at the cell cortex (Komarova et al. 2002), and microtubule stabilisation occurs through the close coupling of rescue and catastrophe events, holding microtubules in a dynamic captured state with short length fluctuations (Straube 2011; Straube and Merdes 2007). Microtubules are captured at the leading edge’s cell cortex by a number of pathways, including EB1/APC/mDia1, LL5beta/ELKS/CLASPs, IQGAP/CLIP-170 and Dlg (Akhmanova et al. 2001; Drabek et al. 2006; Kroboth et al. 2007; Kumar et al. 2009; Nakamura et al. 2001; Pfister et al. 2012; Schober et al. 2009; Watanabe et al. 2009a; Wittmann et al. 2004). Microtubule capture can be maintained for prolonged times resulting in stable microtubules leading to the front of the cell. These long-lived microtubules in turn acquire a number of posttranslational modifications. While detyrosination protects microtubules from depolymerases and severing enzymes (Peris et al. 2009; Roll-Mecak and Vale 2008), acetylation and polyglutamylation recruit microtubule-severing enzymes (Lacroix et al. 2010; Sudo and Baas 2010). Microtubule severing close to the cell cortex can result in the release of a captured microtubule and is a mechanism that allows the spatial regulation of microtubule stability (Zhang et al. 2011).
The inactivation of the microtubule destabilisers stathmin and MCAK at the front of the cell by phosphorylation results in a gradient of increasing microtubule stability towards the front of the cell (Braun et al. 2014; Niethammer et al. 2004). Likewise, the interaction of microtubules with focal adhesion sites results in different outcomes at the front and rear of the cell: While microtubules are captured at adhesion sites in the front of the cell (Kaverina et al. 1998), catastrophe is induced when microtubules contact trailing adhesions (Efimov et al. 2007). While the mechanisms underlying these differences remain to be understood, it is clear that microtubule dynamicity is crucial for cell migration. Freezing dynamicity with low doses of Taxol and other microtubule-targeting agents so that the overall organisation is not perturbed impairs protrusion in fibroblasts, migrating neurons and growing axons (Dunn et al. 1997; Liao et al. 1995; Rochlin et al. 1996; Tanaka et al. 1995; Umeshima et al. 2007; Vasiliev et al. 1970). Furthermore, interference with the dynamicity of rear microtubules specifically leads to decreased rear retraction and changes to the time HeLa cells and CHO fibroblasts spent migrating (Ganguly et al. 2012). When the regional differences in microtubule dynamics regulation are removed by inhibition of MCAK or constitutive activity of Rac1, directional cell migration is severely reduced (Braun et al. 2014).
6.4 How Do Microtubules Influence Cell Migration?
6.4.1 Cell Shape, Polarity and Directionality
Directional cell migration requires the establishment of distinct regions in the cell as the front and the rear. This is often reflected in the morphology of the cell, where the leading edge is protruding either as a flat lamellipodium, using spiky filopodia, pseudopods or more complex structures such as the leading process of neurons. Retracting rears can be either (1) curved inwards pushing against the nucleus as in keratinocytes, (2) long, tail-like extensions as in some epithelial cells and fibroblasts or (3) uropods in leucocytes (Keren et al. 2008; Ratner et al. 1997; Theisen et al. 2012). In each configuration, the protruding edge, the nucleus and the retracting rear set up a single polarity axis. When branches or multiple protrusions are formed, these are often used to make directional decisions in chemotaxis and neuronal pathfinding with the better-positioned protrusion persisting (Andrew and Insall 2007; Cooper 2013). Directional protrusions for cell migration are very similar to emerging axons. In some neurons such as cortical projection neurons, the axon is formed during cell migration by extending cell tails that continue to grow rather than retract (Cooper 2013).
Yet how is the polarity axis established? In cells that have been “starved” by serum withdrawal and then exposed to a chemical attractant gradient, a protruding extension is established towards the higher concentration of the chemical, and the cell begins to move up the gradient. A very similar mechanism guides axon growth cones along attractive or repulsive gradients (Vitriol and Zheng 2012). This mechanism has been conserved from amoeba to humans (Van Haastert and Devreotes 2004). Even in the absence of a guiding chemical gradient, cells from higher eukaryotes that are not surrounded by others spontaneously polarise and form a lamellipodium at one side of the cell. In keratinocytes, symmetry breaking occurs by contraction of actin filaments by non-muscle myosin II on one side of the cell, leaving the opposite side free to protrude (Yam et al. 2007). In epithelial cells, adhesion at the rear and formation of a tail precede protrusion in the opposite direction (Rid et al. 2005; Vicente-Manzanares et al. 2009). Pulling forces from other cells in a collective result in protrusion at the opposite cell edge, resulting in mechanical feedback and coupling of collective cell migration (Weber et al. 2012).
For a cell to change direction, either the polarity axis is gradually shifted, the cell depolarises and repolarises again in a new direction, or the front bifurcates or branches with one of the new protrusions taking over as front after a while (Petrie et al. 2009). The latter mechanism of branching and retraction of a branch is a pathfinding mechanism, for example, in migrating cortical interneurons and neocortical neurons (Cooper 2013; Sakakibara et al. 2014).
How do microtubules support the establishment, maintenance and changes of the polarity axis? As explained above, the asymmetry in the microtubule organisation and distribution of posttranslational modifications enables intracellular trafficking along microtubules to be asymmetric. Important cargo for cell polarity and migration is generated in and near the nucleus in the cell centre and requires transport along microtubules for delivery to the cell edges. An example is the mRNA for β-actin, which localises to the leading edge of migrating cells and is transported by kinesin-1 and dynein along microtubules (Kislauskis et al. 1997; Ma et al. 2011). The localised translation of actin mRNA is important for directional cell migration as it dictates the sites of actin filament nucleation (Katz et al. 2012). Equally importantly, proteins modified and packaged in the Golgi apparatus are transported efficiently to the leading edge via front-directed microtubules nucleated at the trans-Golgi by CLASPs (Miller et al. 2009). Further important cargoes to support front protrusion are vesicles that can be used as a source for additional membrane and supply receptors for adhesion helping protrusion at the leading edge (Etienne-Manneville 2013). It can be beneficial to distribute receptors for sensing chemical gradients and to adhere to the extracellular substrate and neighbouring cells unequally at the cell surface to enhance or adapt to extracellular signals and regulate adhesion in different parts of the cell.
Given that the asymmetry in the microtubule cytoskeleton is key to directional intracellular transport, factors that regulate centrosome positioning such as Lis1 are implicated in developmental diseases due to impaired neuronal migration. Lis1 interacts with dynein to regulate the forces acting on cortical microtubule ends and thereby the centrosome and is crucial to moving the nucleus forward, an essential step in neuronal migration (Umeshima et al. 2007). The loss of Lis1 leads to a smooth brain surface, abnormal neuronal layering and large brain ventricles in humans (Ozmen et al. 2000; Pilz et al. 1998). Similar defects in brain morphology are caused by insufficient neuron migration upon loss of Dcx (doublecortin) (Gleeson et al. 1999a; Gleeson et al. 1999b; Liu 2011; Pilz et al. 1998). Dcx is a MAP that increases microtubule stability, but can also interact with Lis1 (Caspi et al. 2000). Centrosome position also determines the site of axon growth when hippocampal neurons differentiate (de Anda et al. 2005). It is thought that centrosome position again creates a bias of microtubules towards specific sites of the cells, with consequences for intracellular trafficking, protrusion, adhesion and signalling. In line with this idea, the amplification of centrosomes results in increased protrusion and invasion, probably by increasing front-directed microtubule activities (Godinho et al. 2014). For additional information on neurodevelopmental disorders caused by defective cell migration, please also consult the Chap. 5 by Gambarotto and Basto and the Chap. 4 by Sánchez-Huertas, Freixo and Lüders.
It is now firmly established that signalling by small GTPases of the Rho family is important in cell polarity (Nobes and Hall 1999). Small Rho GTPases are proteins that are active in the GTP-bound state, and their activity is regulated by guanine nucleotide exchange factors (GEFs) and GTPase-activating proteins (GAPs). GAPs accelerate GTP hydrolysis to switch off the Rho GTPase, while GEFs accelerate the removal of the product and binding of GTP to activate Rho GTPase signalling. Once activated, Rho GTPases bind to a number of effectors such as protein kinases and actin-binding proteins (Sit and Manser 2011). In migrating cells, the most important Rho GTPases are Rac1, Cdc42 and RhoA. Their activity regulates cell polarity: Rac1 is most important in regulating the protrusion of cells through the WAVE and Arp2/3 complex (Eden et al. 2002). Cdc42 is most active at the cortical zone to promote protrusion via the WASP pathway and is important in orienting the centrosome towards the leading edge via the PAR complex, dynein and microtubules (Etienne-Manneville et al. 2005; Palazzo et al. 2001). RhoA is active further into the lamella and at the rear of the cell to regulate actin contractility (Amano et al. 2010; Machacek et al. 2009). Microtubules are known to influence the activity of Rho GTPases through the local distribution and function of GEFs, GAPs and effectors. Growing microtubules activate Rac1, while the release of microtubule-bound GEF-H1 upon microtubule depolymerisation activates RhoA (Nalbant et al. 2009; Ren et al. 1998); thereby, microtubule dynamics supports the localised activity of Rho GTPases. In turn, GTPases also influence microtubule stability in a positive feedback loop to improve cargo delivery to sites of active protrusion, e.g. RhoA stabilises microtubules via IQGAP1 and mDia1 (Brandt et al. 2007; Kholmanskikh et al. 2006; Wen et al. 2004; Wittmann et al. 2004).
A second connection between microtubules and cell polarity is established through the interaction of microtubules with the Par complex (Suzuki and Ohno 2006). The Par complex, composed of Par6, atypical protein kinase C and Par3, regulates centrosomal polarity. The complex acts downstream of Cdc42 and regulates the activity of GSK3 kinases, which in turn control the activity of MAPs and thereby influence microtubule dynamics locally at the leading edge (Etienne-Manneville et al. 2005). A related protein, MARK/Par-1, can detach MAPs from microtubules to destabilise them (Ebneth et al. 1999; Tassan and Le Goff 2004). MARK activity is highest at the rear and lowest at the front of the cell, increasing the front-biased asymmetry in the microtubule array (Hayashi et al. 2012).
Recently, evidence is accumulating that maintaining an extended cell rear can influence persistent motility. The maintenance of such a tail requires adhesion at the rear despite high contractile forces. Reduction of contractile forces allows formation of extended tails in CHO cells and increases cell motility (Vicente-Manzanares et al. 2007). Likewise, microtubule transport of integrins into cell tails is required for the maturation of trailing focal adhesions and the stability of cell tails. Interfering with microtubule transport by depletion of the kinesin motor Kif1C results in shortened lifetime of cell tails and more frequent directional changes in migrating cells (Theisen et al. 2012). Similarly, drug treatments that suppress dynamic microtubules in the rear of the cell led to increased tail stability and affected directionality in HeLa and CHO cell (Ganguly et al. 2012). In these cells, the morphology of the front of the cells was not affected nor was the front-oriented position of the centrosome, arguing that the cells’ ability to polarise was not globally perturbed. One hypothesis is that drag generated at the cell rear acts as a mechanical cue to support protrusion in the opposite direction (Theisen et al. 2012; Weber et al. 2012). Likewise, the extended cell polarity axis could facilitate biochemical gradients and cytoskeletal filament orientation (Rid et al. 2005; Theisen et al. 2012).
In sum, microtubules have important functions in supporting cell polarity by ensuring that signalling and actin-dependent processes are asymmetric. The interactions with the actin cytoskeleton are likely to function as a positive feedback loop, in which microtubules deliver actin-regulating proteins, while proteins localising to the actin cortex enhance microtubule stability (Siegrist and Doe 2007).
6.4.2 Force Generation
Forces generated by microtubules themselves are generally thought to be of minor importance for moving a cell forward. Microtubules can generate pushing and pulling forces through coupling polymer assembly and/or disassembly to subcellular structures. These forces are harnessed in the movement of chromosomes during mitosis and contribute to the distribution of the endoplasmic reticulum (Jordan and Wilson 2004; Waterman-Storer and Salmon 1998). In cell types where only a small number of pioneer microtubules reach the plasma membrane, the direct contribution of microtubules to membrane protrusion is probably not significant. However, large numbers of microtubule ends reach the cell edge in axonal growth cones, and pushing forces generated by assembling microtubules are likely to be harnessed for cell protrusion (Liu et al. 2010). In this system, microtubule motors also generate forces either by sliding two microtubules relative to each other or by moving microtubules relative to the cell cortex, so that more microtubule ends reach the cortex. The main motors implicated in microtubule motility are kinesin-1 and dynein. Kinesin-mediated microtubule-microtubule sliding has been shown to generate forces for the protrusion of neurites (Lu et al. 2013; Myers and Baas 2007). To which extent forces generated by microtubule sliding and polymerisation directly contribute to cell migration remains to be established as microtubules also affect cell protrusion by a number of indirect pathways, most of which involve the actin cytoskeleton. It is well accepted that pushing forces generated by the assembly of actin at the cell front are the main driving force for cell protrusions. Likewise myosin-mediated contraction of actin bundles generates hydrostatic pressure and contractile forces involved in protrusion as well as contraction. Therefore, force generation during cell migration is primarily attributed to the action of the actin cytoskeleton.
Microtubules support actin-mediated cell protrusion indirectly through delivery of vesicles, i.e. lipids to the cell front, thereby allowing the expansion of the plasma membrane at the leading edge. The positioning of mRNA for actin and Arp2/3 at the leading edge is likely to involve microtubule-based transport and ensures a ready supply of actin monomers and the main actin nucleator for lamellipodial protrusion at the front of the cell (Jaulin and Kreitzer 2010; Mingle et al. 2005; Oleynikov and Singer 1998). Furthermore, the microtubule plus end complex contains a number of actin nucleators and regulators. Amongst them is adenomatous polyposis coli (APC), a protein that also promotes microtubule assembly (Kita et al. 2006; Mimori-Kiyosue et al. 2000) and acts as an actin nucleator in synergy with the formin mDia1 (Nathke et al. 1996; Okada et al. 2010). In addition to APC, a number of MAPs have been identified to bind and/or regulate both microtubules and actin. These include CLASPs, ACF7, MAP4 and dynein/dynactin (Matsushima et al. 2012; Rodriguez et al. 2003; Tsvetkov et al. 2007; Wu et al. 2008). For example, GSK3β acts downstream of the polarity-regulating GTPase Cdc42 and controls microtubule stability via ACF7 and other factors (Etienne-Manneville and Hall 2003; Kodama et al. 2003). ACF7 itself cross-links actin and microtubules, influences microtubule dynamics and has microtubule guidance functions (Applewhite et al. 2010; Wu et al. 2008). Also the non-receptor tyrosine kinase ABL2/Arg binds to microtubules and actin and promotes cell protrusion and spreading. This activity requires the physical coupling between F-actin and microtubules by ABL2 (Miller et al. 2004).
Other ways in which microtubules can influence actin polymerisation are by locally regulating small GTPase signalling, which in turn regulate force generation. It has been known for some time that microtubule polymerisation can activate Rac1 (Montenegro-Venegas et al. 2010; Waterman-Storer et al. 1999). Microtubules bind the Rac1 activators Tiam1, Stef and Trio (Pegtel et al. 2007; Rooney et al. 2010; van Haren et al. 2014), thus allowing microtubule-dependent regulation of Rac1 through several pathways. RhoA can be activated by GEF-H1, which is sequestered on the microtubule lattice and activated upon release during microtubule catastrophe (Nalbant et al. 2009; Ren et al. 1998).
Thus a complex network of structural and signalling interactions between the microtubule and actin cytoskeleton at the cell front controls cell migration, and a fine balance between these activities is important for robust and directional cell migration (Kaverina and Straube 2011). So far, no diseases have been linked to an imbalance of forces in cell migration, but as many of the players involved serve multiple functions, and we do not yet fully understand how they interact with each other, it may be possible that we are underestimating the significance of a force imbalance for disease development. This intriguing area awaits further investigation, but individual players (e.g. APC, RASSF1A) have already been demonstrated to play important roles in cancer development (Humbert et al. 2008; Kassler et al. 2012; van Es et al. 2001).
While actin and non-muscle myosin II provide the forces necessary for protrusion at the leading edge, the microtubule cytoskeleton with its motor dynein can supplement these forces when necessary. In elongated cells that need to move in coherent clusters within surrounding tissue pressing in on them, such as migrating neurons, moving the nucleus presents a difficult challenge (Harada et al. 2014). The nucleus is the bulkiest organelle in the cell that cannot easily be compressed without causing DNA damage. Hence moving it against pressure from the environment requires forces that exceed those that actin rear contraction can provide (Tsai et al. 2007). The close spatial localisation of the centrosome to the nucleus in interphase cells has suggested early on that microtubules might be important in this task. Experiments on granule cells from mice explant cultures could demonstrate that stable microtubules and dynein are essential to move the nucleus and to position the centrosome in front of the nucleus (Tsai and Gleeson 2005; Umeshima et al. 2007). These results have led to two models on how microtubules and dynein can be used to move the nucleus (nucleokinesis): One model suggests that dynein is anchored at the leading edge to pull on plus ends of microtubules whose minus ends are embedded in the centrosome, which serves to translate the forces from dynein into net forward movement of the nucleus (Tsai and Gleeson 2005). Another model implicates a cage formed from a subpopulation of acetylated microtubules that encloses the nucleus and transmits the force generated by cortex-anchored dynein to move the nucleus forward (Umeshima et al. 2007). It should be noted though that not all neurons use dynein-mediated forces to move their nuclei. Differences exist between types of neurons and between the same neuron types in different organisms. For example, different force-generation models implicating actin-generated pushing forces exist for cerebellar Purkinje cells and cortical interneurons and also for cerebellar granule cells from mice and zebrafish (Cooper 2013). One possible explanation for these differences was proposed to lie in the different cell shapes, as the wider zebrafish cells might be able to move the nucleus by actin-mediated contractility alone, while the very narrow and elongated mouse neurons require additional microtubule-mediated forces (Cooper 2013).
In order for the cell to move forward, the forces generated through actin polymerisation and contraction need to be transmitted to the extracellular matrix or neighbouring cells. To achieve this, cells form adhesive structures: focal adhesions and podosomes that attach to the extracellular matrix, and tight junctions, gap junctions and adherens junctions that link them to neighbouring cells. The size and composition of these structures depend on the type of cell and the cellular environment. Typically, adhesive structures are formed by a transmembrane receptor, which contacts the substrate on the outside of the cell or forms homophilic interactions with the neighbouring cells. The receptor is then stabilised on the inside of the cell by association with other proteins. The adhesion complexes are connected to the cytoskeleton, which will also contribute to clustering of such complexes into larger structures.
The dependence of cells on adhesion for migration can be very different. In confined environments, protrusions such as blebs can generate enough traction themselves to allow the cell to move forward efficiently. Pressurised blebs can be used to find the weakest linkage between cells and can create a foothold for moving cells trying to cross tissues (Lammermann and Sixt 2009; Mandeville et al. 1997; Sanz-Moreno and Marshall 2010; Wolf et al. 2003b). Such modes of migration are employed by cells of the immune system, such as neutrophils and leucocytes, and some tumour cells (Friedl et al. 1998a; Friedl et al. 1998b; Werr et al. 1998). Mesenchymal migration of fibroblasts and epithelial cells relies strongly on cell adhesion for migration in 2D as well as in 3D (Sanz-Moreno and Marshall 2010). Adherent cells can use different classes of receptors to attach to their surroundings; the classic receptors for a variety of extracellular matrix molecules are integrins. Integrins are obligatory heterodimers of an α- and a β-chain, and different combinations of the 18 α- and 8 β-chains in mammalian cells result in 24 different receptors with distinct substrate specificity (Hynes 2002). Integrins are embedded in the plasma membrane with the greater part of the protein extending into the extracellular space where it directly binds to matrix proteins. Exocytosis of integrin-containing vesicles delivered by microtubule-dependent trafficking occurs at the leading edge (Bretscher and Aguado-Velasco 1998; Spiczka and Yeaman 2008) allowing the formation of small focal complexes. At least in part, this process is controlled by Rac1 which becomes activated by Tiam2, which in turn is regulated by microtubules (Rooney et al. 2010). Focal complexes turn over rapidly with only a few of them maturing into focal adhesions. Focal adhesions consist of >150 proteins on the cytoplasmic side, which mediate links to actin fibres and/or function in signalling (Zaidel-Bar et al. 2007). Focal adhesion maturation is force dependent: Actin contractility increases the size of adhesions as well as the density of adhesion molecules in the adhesion (Parsons et al. 2010). This response allows adhesion strength to scale to the forces applied to them.
Microtubules are important regulators of focal adhesions. The disassembly of microtubules by small-molecule inhibitors results in the formation of large focal adhesions, while their turnover is induced as soon as microtubule regrowth is permitted by washing out of the drug (Ezratty et al. 2005; Waterman-Storer et al. 1999). Furthermore, microtubules have been observed to target focal adhesions repeatedly with their dynamic plus ends, and this targeting results in the dissolution of focal adhesions (Kaverina et al. 1999; Kaverina et al. 1998; Krylyshkina et al. 2003; Rid et al. 2005). Microtubules are thought to reach focal adhesions by guidance along actin filaments. In migrating fibroblasts, microtubules are crossbridged to actin filaments by a number of factors including ACF7, IQGAP1/CLIP-170 or CLASPs, which then guide the growing microtubule ends to focal adhesions (Drabek et al. 2006; Small and Kaverina 2003; Stehbens and Wittmann 2012). Microtubule ends reduce their growth speed and undergo catastrophe upon contact with focal adhesions. This process is regulated by paxillin, a structural component of focal adhesions (Efimov et al. 2008). Often, the microtubule undergoes a rescue and targets the same or another focal adhesion, thereby resulting in the repeated targeting of adhesions and their turnover.
One possible way how microtubules could disassemble focal adhesions is by interacting with signalling molecules that control the composition of focal adhesions (Etienne-Manneville 2013; Wickstrom et al. 2010), and another is that microtubules deliver components of the endocytic machinery, as could be shown for dynamin and Clathrin, to help internalise integrins for recycling (Chao and Kunz 2009; Ezratty et al. 2009; Nishimura and Kaibuchi 2007). Also, microtubule-dependent control of the local release of proteases into the extracellular space may promote the detachment of the cell from the substrate by cleaving substrate-bound receptors (Takino et al. 2006). It was demonstrated that exocytosis of such proteases occurs in the vicinity of focal adhesions (Steffen et al. 2008; Wiesner et al. 2010), but if this mechanism plays a role in cell migration remains to be established (Margadant et al. 2011). It is, however, well known that localised secretion of metalloproteases is important for the migration of cancer cells through existing tissue (Hegerfeldt et al. 2002; Takino et al. 2006; Wang and McNiven 2012; Yilmaz and Christofori 2009). Blocking these proteases stops the migration of fibrosarcoma and mammary carcinoma cells (Coopman et al. 1998; Wolf et al. 2003a). Likewise, microtubule-dependent regulation of actin dynamics (see section above) could affect the force coupling into focal adhesions with loss of the pulling force resulting in the dissolution of the focal adhesion.
The microtubule-dependent control of focal adhesions requires motor-dependent transport as kinesin-1 has been demonstrated to be required for the process (Krylyshkina et al. 2002). Podosomes, invasive adhesion structures prevalent in immune cells such as macrophages and dendritic cells, require the kinesin-3 Kif1C for their formation and dynamic turnover and Kif9 for their function in matrix degradation via localised exocytosis (Bachmann and Straube 2015; Cornfine et al. 2011; Efimova et al. 2014; Kopp et al. 2006). However, it is currently not clear which cargoes are delivered by these kinesins that contribute to the observed processes.
Controlled turnover of focal adhesions is likely to play a role in the metastatic behaviour of cancers, regulating the aggressiveness of disease progression by the cells’ motility and invasiveness (McLean et al. 2005; Recher et al. 2004). The formation of adhesions is in the range of several minutes, which can be the rate-limiting step in migration as shown by the increase in cell migration speed in vinculin-depleted cells (Friedl et al. 2004; Mierke et al. 2010). In accordance with this, a reduction in cell adhesiveness has been implicated in the progression of cancer (Sanz-Moreno and Marshall 2010). Cells migrating as collective, either as clusters of cancer cells or during developmental processes, need to maintain close connections to the other cells at all times in order to improve their migration efficiency, as surrounding tissues pose significant obstacles. Cadherins play an important role in this.
Cadherins are a large family of membrane-bound receptors that form homophilic interactions with molecules on the surface of neighbouring cells. This establishes a tight link between cells. Examples of cells that depend on N-cadherin for motility are a number of different types of migrating neurons (Jossin and Cooper 2011; Lele et al. 2002; Monier-Gavelle and Duband 1995; Nakagawa and Takeichi 1998; Rappl et al. 2008; Rieger et al. 2009) but also cells forming the lateral line organ in zebrafish (Revenu et al. 2014) and cancer cells (Qi et al. 2006; Shih and Yamada 2012). Other cells rely on E-cadherin, such as fibroblasts and keratinocytes (Maretzky et al. 2005). The increased cohesion mediated by cadherin within the cell cluster could facilitate pulling of follower cells along the path that the leader cells have created by breaking down the extracellular matrix (Friedl and Gilmour 2009), or it could provide a point of strong attachment for cytoskeletal elements to help move cell organelles like the nucleus forward, especially in neurons (Rieger et al. 2009; Tsai and Gleeson 2005). Like most other plasma membrane-bound proteins, cadherins require kinesin-based transport to reach their destination (Chen et al. 2003; Kawauchi et al. 2010; Mary et al. 2002; Yanagisawa et al. 2004). In addition, the plus ends of non-acetylated microtubules have been shown to cluster cadherins in the plasma membrane, a prerequisite to forming stable cell-cell connections (Stehbens et al. 2006; Waterman-Storer et al. 2000). Similar to cadherins, CAMs are a large group of proteins that can form homophilic interactions to connect two cells. They are often upregulated when cells obtain increased motile characteristics such as during metastasis (Lehembre et al. 2008; Schreiber et al. 2008). They possess functions in addition to adhesion, such as sensing chemical gradients during migration, making their regulation even more complex (Cavallaro et al. 2001; Francavilla et al. 2007; Paratcha et al. 2003; Yilmaz and Christofori 2009).
All these different types of adhesions have their own signalling pathways, which link adhesions and their various states of engagement to polarity signalling and microtubule stability, and they all depend on microtubule-based transport from the cell centre to the surface. This places microtubule-mediated transport at the centre of the regulation of local adhesiveness by site-directed delivery of substrate receptors or regulatory elements (Miller et al. 2009; Yadav et al. 2009). Many cell surface proteins have residency times at the surface in the range of seconds to minutes (Bretscher 2008), before they need to be internalised and either transported back into the cells for processing or returned to specific sites to counteract diffusion in the plasma membrane. For N-cadherin and α5β1 integrin, for example, recycling pathways have been described which can be rather elaborated, involving internalisation, retrograde transport to recycling compartments that can be as far away as next to the centrosome and return to the surface (Bretscher 1989; Caswell and Norman 2008; Gu et al. 2011; Shieh et al. 2011). Through their transport capacity and motor protein preference for specific microtubule tracks, cargo can be directed to different parts of the cell (Cai et al. 2009; Reed et al. 2006), giving microtubules control over the amount and position of adhesive complexes on the cell surface. For example, Kif1C transports integrin-containing vesicles in migrating cells. This transport is required for the maturation of focal adhesions in the rear of the cell as it provides the ready supply of integrins for additional incorporation and exchange. A reduced supply of surface integrin results in a misbalance of contractile forces and adhesion strength causing the frequent contraction of cell tails and loss of polarity (Theisen et al. 2012). Recently, kinesins Kif15 and Kif4A have also been implicated in integrin transport (Eskova et al. 2014; Heintz et al. 2014). How the different transport pathways contribute to the microtubule-dependent regulation of cell adhesion remains to be elucidated.
The coordination of the cell migration machinery at the front and rear of the cell and the response to environmental signals and guidance cues involve complex signalling networks. Amongst the well-characterised pathways organising migration are polarity signalling (small GTPases), adhesion signalling (integrins and cadherin) and guidance signalling (with the use of second messengers, intracellular calcium and phosphoinositol species).
Rho GTPases regulate actin dynamics, contractility and cell adhesion (Sit and Manser 2011). Rho GTPase signalling pathways are spatially restricted allowing the local regulation of protrusion and retraction enabling cell migration and other processes such as cytokinesis, phagocytosis and morphogenesis (Hall 2012). Microtubules control Rho GTPases signalling (1) by delivery of GTPases Rac1 and Cdc42 to the membrane (Osmani et al. 2010; Palamidessi et al. 2008); (2) by positioning GEFs such as Tiam1, Stef/Tiam2, Trio and effectors such as IQGAP1 (Briggs et al. 2002; Briggs and Sacks 2003b; Rooney et al. 2010; van Haren et al. 2014); and (3) by sequestering GEFs and coupling their release and activation to microtubule dynamics such as GEF-H1/RhoGEF2 (Chang et al. 2008; Glaven et al. 1999; Krendel et al. 2002; Rogers et al. 2004). In turn, Rho GTPases regulate microtubule dynamics. In cells without the Rac1 GEF Tiam1, microtubules are unstable (Pegtel et al. 2007), and Cdc42 influences the polarity of the microtubule array via the Par complex and GSK3β (Etienne-Manneville et al. 2005; Watanabe et al. 2009a). Therefore, the relationship between microtubules and GTPases is balanced by feedback loops (for further examples, see review by Etienne-Manneville 2013).
Rho GTPase signalling is connected to adhesion signalling. Cadherins at the plasma membrane are signalling hubs via their binding of β-catenin and p120. β-catenin can be released from cadherin to move into the nucleus and, as co-factor, triggers the transcription of several genes, including those of adhesion molecules (McCrea et al. 2009). This is a crucial event in Wnt signalling, a pathway that is often enhanced in cancer cells and metastasis and which is controlled by Cdc42 (Aman and Piotrowski 2008; Clevers 2006; Fukata et al. 1999; Heuberger and Birchmeier 2010). Another component of the Wnt signalling pathway is APC, which is localised at the leading edge at microtubule plus ends (Matsumoto et al. 2010; Okada et al. 2010) and which regulates β-catenin levels (Munemitsu et al. 1995). The release of β-catenin from cadherin is also partly depending on IQGAP1, which is an effector of Rho GTPases and can bind microtubules directly to stabilise them (Fukata et al. 1999; Fukata et al. 2002). p120, another catenin family protein normally found associated with cadherin, has been reported to suppress RhoA and increase the activity of Rac1 and Cdc42 to regulate cell-cell contacts and may be able to influence microtubule dynamics (Ichii and Takeichi 2007; Watanabe et al. 2009b).
Another example for crosstalk between polarity signalling and adhesion signalling is the relationship between small GTPases and integrin. Integrin signalling is activated by binding of integrins to the extracellular substrate and is mostly mediated through focal adhesion kinase (FAK) and integrin-linked kinase (ILK) (Schwartz 2001). FAK regulates the turnover of focal adhesions but also activates RhoA and mDia (Palazzo et al. 2004; Webb et al. 2004). As mDia can bind to microtubule plus ends at the leading edge, this could explain the observed link between FAK activity and microtubule stabilisation (Palazzo et al. 2004). Focal adhesions can also influence the activity of Cdc42, which can act back on microtubule stability (Etienne-Manneville and Hall 2001). In migrating neurons, interfering with the function of FAK leads to a disorganised microtubule array and defective nuclear movement, a prerequisite for neuronal migration (Xie et al. 2003). Similarly, ILK regulates Rac1 and therefore lamellipodium formation via its interaction partners α- and β-parvin (Legate et al. 2006; Zhang et al. 2004). ILK and microtubules together function to impart polarity on epithelial cells, and ILK is needed to organise microtubules in this system (Akhtar and Streuli 2013). Other effects of ILK include the regulation of microtubule dynamics through the interaction with IQGAP1 and mDia1 (Wickstrom et al. 2010).
Recently, it was proposed that local intracellular calcium levels, a second messenger common to many signalling pathways, could be another mechanism to coordinate the different signalling pathways and biological processes (Tsai et al. 2014). Calcium waves at the front of migrating fibroblasts dictate cell speed. As some of the microtubule-regulating proteins such as IQGAP1 require calmodulin and/or calcium for their function (Briggs and Sacks 2003a), it is possible that other signalling pathways which we currently do not know can influence microtubules by these means indirectly and thereby increase the microtubule-centred regulatory network during migration.
While many cell types are able to generate movement in the absence of microtubules by employing their actin cytoskeleton, microtubules are consistently important in fine tuning several aspects of migration, such as establishing polarity, exercising spatial control over force generation and adhesion, as well as signalling. Microtubules span the entire cell, making it possible to coordinate these tasks across spatially distant cellular regions. Due to their intrinsic dynamic instability, microtubules can adapt quickly in response to external and internal cues.
Over recent years, it has become clear that imbalance or mis-regulation of microtubule dynamics and/or motor function can lead to disease or promote disease progression when cells that should move cannot (e.g. immune cells or cells in embryonic development) or cells that should not move gain the ability to break down tissue barriers and colonise other tissues (e.g. cancer metastasis). Further research will continue to elucidate the details of the molecular interactions and will help us to understand the development of diseases affecting many patients.
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