Microtubules Regulate Cell Migration and Neuronal Pathfinding

Chapter

Abstract

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.

6.1 Introduction

Cell migration is a fundamental biological phenomenon occurring in protists as well as in multicellular organisms. Locomotion of unicellular organisms enables access to nutrients and optimal environmental conditions as well as the assembly of cells into spore-bearing structures (Van Haastert and Devreotes 2004). In multicellular organisms, migration is essential to positioning each cell in the body at its correct location. During embryonic development, many cells are generated from precursors in a different location to where they are needed. Furthermore, neuronal precursors in mammals need to migrate not only to reach a specific destination but also to encounter the correct type of cells along the way to form contacts with in order to build the neuronal network in the brain. For example, cerebellar granule cell precursors migrate tangentially until they change to radial migration along glial fibres during which they establish contacts with Purkinje cells needed for the proper wiring of the adult cerebellum (Cooper 2013; Komuro and Rakic 1998; Fig. 6.1a). Other instances of migration occurring during development are clusters of cells that move along the entire length of the body to form the lateral line organ in fish (Fig. 6.1b) and precursors of muscle cells that align before fusion into muscle fibres (Revenu et al. 2014; Wakelam 1985). In adults, cell migration is of utmost importance for immune surveillance and response and for healing wounds (Fig. 6.1c). Finally, defective regulation of migration contributes to chronic inflammatory diseases such as gout and atherosclerosis and enables the spreading of cancer cells from the primary tumour site (Chi and Melendez 2007; Colvin et al. 2010; Friedl and Wolf 2003, Fig. 6.1d). Metastasis is responsible for 90 % of cancer deaths (Mehlen and Puisieux 2006). For an overview of human diseases directly linked to defective migration as a result of an impaired microtubule cytoskeleton, please refer to Table 6.1.
Fig. 6.1

Examples of migration modes and cell shapes. (a) Modes of migration depend on the cellular environment. In this example, a cerebellar granule cell neuron (white) is migrating tangentially over other tissue, until it finds a glia cell (dark grey). It is then guided along the axon of the glia cell during radial migration using microtubule-dependent nucleokinesis to reach the inner layer of the developing cerebellum. During radial migration, the axon of the granule cell projects from the rear of the cell and establishes contacts to Purkinje cells (light grey) (Fahrion et al. 2012). (b) Some cells migrate as highly coordinated multicellular strands during development. In this example, leader cells form a path for the follower cells. In the lateral line primordium, these take the form of rosettes. Close communication between leader and follower cells is necessary to achieve collective migration (Revenu et al. 2014). (c) Similarly, cancer cells often metastasize as clusters of cells that force their way by localised release of extracellular matrix-degrading enzymes (Friedl and Gilmour 2009). (d) Wound healing can be recreated in cell culture. Typically, once a gap is created, individual cells (leader cells, light grey) at the edge of the wound sense the gap and respond by extending a lamellipodium into the gap. Once these cells start to invade the open space, they are followed by other cells pushing from behind (Tsai et al. 2014). (e) Commonly observed migrating cell types are depicted in their relative size. All cells migrate towards the top of the page. Note that the cell area depends on the mode of migration and the stiffness of the substrate and can therefore change depending on the cellular environment

Table 6.1

Diseases directly associated with cell migration and microtubule function

Cause/accelerating factor

Effect

Disease

Reference

Microtubule structure and stability:

   

Dynamic microtubules

Increased motility

Metastasis

Mehlen and Puisieux (2006)

Dynamic microtubules

Immune cell migration

Rheumatoid arthritis

Brahn et al. (1994), Friedl and Weigelin (2008)

Dynamic microtubules

Neutrophil migration

Gout

Chia et al. (2008)

Dynamic microtubules

Infiltration of immune cells into the brain; multiple sclerosis-like phenotype

Experimental autoimmune encephalomyelitis

O’Sullivan et al. (2013)

TUBB3

Neuronal migration defects

Malformation of cortical development

Poirier et al. (2010), Saillour et al. (2014)

TUBA1A

Neuronal migration defects

Lissencephaly/pachygyria

Poirier et al. (2007)

TUBB2B

Neuronal migration defects

Polymicrogyria

Jaglin et al. (2009)

TUBB5

Neuronal migration defects

Microcephaly

Breuss et al. (2012)

TUBA3A

Neuronal migration defects

Polymicrogyria

Keays et al. (2007)

TUBG1

Neuronal migration defects

Malformation of cortical development

Poirier et al. (2013)

Doublecortin

Neuronal migration defects

Lissencephaly

Gleeson et al. (1999b), Pilz et al. (1998)

MAP1B

Neuronal migration defects

Diverse neuropathologies

Del Rio et al. (2004), Riederer (2007)

APC

Impaired neuronal network formation

Schizophrenia, autism

Cui et al. (2005), Kozlovsky et al. (2002), Mohn et al. (2014)

HDAC6

Blood vessel formation

Tumour angiogenesis, metastasis

Li et al. (2011), Wu et al. (2010)

Clip-170

Increased vessel density in tumours

Tumour angiogenesis

Sun et al. (2013)

Microtubule length and array control:

   

Tau

Increased microtubule severing

Alzheimer’s disease

Sapir et al. (2012)

Katanin

Sperm motility defective

Male fertility defect

O’Donnell et al. (2012)

Spastin

Impaired microtubule severing

Metastasis

Draberova et al. (2011)

Kif2A

Enhanced cell motility and invasiveness

Metastasis

Wang et al. (2010), (2014)

Motor proteins and their regulation:

   

Dynein

Movement of organelles, retrograde trafficking affected

Charcot-Marie-Tooth disease type 2; several neurological symptoms

Willemsen et al. (2012)

Lis1

Impaired dynein function

Lissencephaly, Miller-Dieker syndrome

Pilz et al. (1998), Badano et al. (2005), Hattori et al. (1994)

Kif5C

Neuronal migration defects

Malformation of cortical development

Poirier et al. (2013)

Centrosome:

   

DISC1

Centrosomal function impaired

Schizophrenia, depression, bipolar disorder

Duan et al. (2007), Hashimoto et al. (2006), Hennah et al. (2009), Ishizuka et al. (2011), Meyer and Morris (2009), Steinecke et al. (2012)

PCM1

Centrosomal satellites defective

Schizophrenia

Kamiya et al. (2008)

SDCCAG8

Centrosomal satellites defective

Schizophrenia

Hamshere et al. (2013), Insolera et al. (2014)

BBS1, BBS4

Defective cilia; defects in migration cause craniofacial dysmorphia

Bardet-Biedl syndrome

Tobin et al. (2008)

Excess centrosomes in interphase

Impaired migration

Angiogenesis; defective vessel sprouting

Kushner et al. (2014)

Cell polarity signalling:

   

Cdc42

Stability of microtubules in the uropod of neutrophils

Immunodeficiency

Kumar et al. (2012)

Cell adhesion:

   

ACF7

Microtubule-regulated adhesion turnover defective

Delayed skin healing

Wu et al. (2008)

APC

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).

In general, cells need to coordinate the following steps in order to achieve migration (Etienne-Manneville 2013; Ridley et al. 2003):
  1. 1.

    Protrusion. This involves the cell membrane to be pushed forward by cytoskeletal polymerisation.

     
  2. 2.

    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).

     
  3. 3.

    Contraction. Actin and myosin generate contractile forces to move the cell body forward.

     
  4. 4.

    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).

By their reach throughout the whole cell, microtubules can coordinate the different aspects involved in cell migration by acting at different parts of the cell at the same time (Fig. 6.2), such as regulating increased adhesiveness at the cell front while reducing adhesiveness at the rear. They are also crucial to long-distance transport and directing cargo (vesicles, proteins, mRNA) to different regions of the cell, thereby gaining a regulatory influence over local protrusion and adhesiveness, signal perception/transduction and cell-cell communication. In addition, their mechanical properties contribute to shaping the cell, for example, by preventing the collapse of membrane structures due to their resistance to compression. Still, the microtubule system is fairly short-lived, as a result of the intrinsic dynamic instability, allowing the microtubule cytoskeleton to adapt very quickly to changes, for example, when signals from the environment are perceived that make changes to the migration direction necessary. In spite of their normally short lifetime, certain microtubules can be stabilised for specific functions, e.g. in order to move the nucleus forward during neuronal migration. Finally, by selectively adapting the composition of proteins binding to the dynamic plus ends, these can provide a spatially and temporally highly restricted environment to carry out special tasks, such as targeting focal adhesions at the rear of the cell for disassembly or interacting with signalling components in a very controlled manner.
Fig. 6.2

Microtubule arrangement in migrating/protruding cells. (a) In epithelial cells moving over a flat surface, more microtubules reach the leading edge than the cell rear. The centrosome nucleates a radial array of microtubules, but rearwards growing microtubules are deflected by the nucleus. In addition, the trans-Golgi nucleates a front-directed microtubule array. This front bias is enhanced by a gradient of microtubule-destabilising factors, which are more active at the rear of the cell and the selective stabilisation of microtubules at the leading edge mediated by plus end capture at the cell cortex. CLASP proteins have been implicated in both nucleation at the Golgi and capture at the cell cortex. (b) In migrating neurons, most microtubules extend towards the leading edge, and only few reach around the nucleus to the rear. Microtubules are nucleated from the centrosome, which is oriented towards the leading edge. A cage of stable microtubules links the centrosome and the nucleus. This cage is important for moving the nucleus forward. (c) Growing axons resemble migrating cells in many aspects. They typically exhibit a dense array of stable microtubules. Microtubules nucleated at the centrosome are often not long enough to reach the leading edge. Instead the array mainly contains free microtubules generated by microtubule severing and capping of the minus ends by stabilising complexes. These microtubules can be moved by motor proteins and contribute to force generation

6.2 Microtubule Organisation in Migrating Cells

In many migrating cells, microtubules show an asymmetric arrangement. This is typically biased towards the front of the cell in most cell types, such as fibroblasts, epithelial and endothelial cells, astrocytes and neurons (Fig. 6.2a, b), but a bias to the rear of the cell has been shown in leucocytes (Kaverina and Straube 2011; Watanabe et al. 2004; Yoo et al. 2012). Many microtubules are nucleated by and anchored with their minus ends at the centrosome so that their dynamic plus ends project towards the cell cortex. Often the centrosome is positioned between the nucleus and the leading edge of the cell. The mechanism behind orienting the centrosome involves microtubule capture at the cortex, which allows the minus-end-directed motor dynein to exert pulling forces on the microtubules to position the centrosome in the cell centre (Palazzo et al. 2001; Tsai and Gleeson 2005; Yvon et al. 2002; Fig. 6.3). In addition, actin-mediated forces pull the nucleus backwards (Gomes et al. 2005). While defects in the positioning of the centrosome are indicative of problems in cell polarity and correlate with cell migration defects (Etienne-Manneville and Hall 2003; Luxton and Gundersen 2011; Tsai et al. 2007), it is unlikely that the position of the centrosome itself determines directionality of cell migration or the asymmetry of the microtubule network. Centrosome position is dictated by cell geometry and therefore a read-out of cell shape (Dupin et al. 2009; Gomes et al. 2005). Most cells moving on 2D surfaces will position the centrosome in front of the nucleus. However, plating the same cells on patterned substrates that confine adhesion to narrow lines results in an elongated cell morphology and efficient cell migration, but the centrosome is found behind the nucleus (Pouthas et al. 2008). Under these conditions, the majority of microtubules still grow towards the front of the cells; therefore, centrosome position and microtubule network bias are independent of each other (Straube, unpublished data). Similarly, in wound-edge Ptk cells, the rear-oriented position of the centrosome may become compensated for by actin-based transport of microtubules to the front (Yvon et al. 2002). Likewise, zebrafish neutrophils migrating in vivo position the centrosome in front of the nucleus, but the majority of microtubules project towards the rear (Yoo et al. 2012) (for an overview of centrosomal positions in different systems, see Luxton and Gundersen 2011). The mechanisms underlying the reverse microtubule orientation in these cells are not understood, and we will concentrate on the more commonly observed front-biased microtubule organisation in the remainder of this chapter.
Fig. 6.3

Microtubule functions in cell migration. Microtubules are the main tracks for intracellular long-distance transport, delivering cargo to support and regulate the cell migration machinery. Microtubules are organised asymmetrically and their stability is regulated spatially by rescue factors such as CLASPs, minus end capping and severing proteins. Dynamic microtubules modulate Rho GTPase signalling by sequestering, concentrating and releasing regulatory proteins. Microtubules stimulate actin polymerisation through delivery of mRNAs and the accumulation of actin nucleators such as APC and formins at microtubule ends. Localised exocytosis supplies the membrane for protrusion and receptors and enzymes for matrix degradation. Microtubule targeting and directed transport also regulates focal adhesions

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).

6.4.3 Adhesion

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.

6.4.4 Signalling

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.

6.5 Conclusion

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.

References

  1. Abal M, Piel M, Bouckson-Castaing V, Mogensen M, Sibarita JB, Bornens M (2002) Microtubule release from the centrosome in migrating cells. J Cell Biol 159:731–737PubMedPubMedCentralCrossRefGoogle Scholar
  2. Akhmanova A, Hoogenraad CC, Drabek K, Stepanova T, Dortland B, Verkerk T, Vermeulen W, Burgering BM, De Zeeuw CI, Grosveld F, Galjart N (2001) Clasps are CLIP-115 and −170 associating proteins involved in the regional regulation of microtubule dynamics in motile fibroblasts. Cell 104:923–935PubMedCrossRefGoogle Scholar
  3. Akhtar N, Streuli CH (2013) An integrin-ILK-microtubule network orients cell polarity and lumen formation in glandular epithelium. Nat Cell Biol 15:17–27PubMedPubMedCentralCrossRefGoogle Scholar
  4. Aman A, Piotrowski T (2008) Wnt/beta-catenin and Fgf signaling control collective cell migration by restricting chemokine receptor expression. Dev Cell 15:749–761PubMedCrossRefGoogle Scholar
  5. Amano M, Nakayama M, Kaibuchi K (2010) Rho-kinase/ROCK: a key regulator of the cytoskeleton and cell polarity. Cytoskeleton 67:545–554PubMedPubMedCentralCrossRefGoogle Scholar
  6. Andrew N, Insall RH (2007) Chemotaxis in shallow gradients is mediated independently of PtdIns 3-kinase by biased choices between random protrusions. Nat Cell Biol 9:193–200PubMedCrossRefGoogle Scholar
  7. Applewhite DA, Grode KD, Keller D, Zadeh AD, Slep KC, Rogers SL (2010) The spectraplakin Short stop is an actin-microtubule cross-linker that contributes to organization of the microtubule network. Mol Biol Cell 21:1714–1724PubMedPubMedCentralCrossRefGoogle Scholar
  8. Bachmann A, Straube A (2015) Kinesins in cell migration. Biochem Soc Trans 43:79–83PubMedCrossRefGoogle Scholar
  9. Badano JL, Teslovich TM, Katsanis N (2005) The centrosome in human genetic disease. Nat Rev Genet 6:194–205PubMedCrossRefGoogle Scholar
  10. Bhuwania R, Castro-Castro A, Linder S (2014) Microtubule acetylation regulates dynamics of KIF1C-powered vesicles and contact of microtubule plus ends with podosomes. Eur J Cell Biol 93(10–12):424–437PubMedCrossRefGoogle Scholar
  11. Brahn E, Tang C, Banquerigo ML (1994) Regression of collagen-induced arthritis with taxol, a microtubule stabilizer. Arthritis Rheum 37:839–845PubMedCrossRefGoogle Scholar
  12. Brandt DT, Marion S, Griffiths G, Watanabe T, Kaibuchi K, Grosse R (2007) Dia1 and IQGAP1 interact in cell migration and phagocytic cup formation. J Cell Biol 178:193–200PubMedPubMedCentralCrossRefGoogle Scholar
  13. Braun A, Dang K, Buslig F, Baird MA, Davidson MW, Waterman CM, Myers KA (2014) Rac1 and Aurora A regulate MCAK to polarize microtubule growth in migrating endothelial cells. J Cell Biol 206:97–112PubMedPubMedCentralCrossRefGoogle Scholar
  14. Bretscher MS (1989) Endocytosis and recycling of the fibronectin receptor in CHO cells. EMBO J 8:1341–1348PubMedPubMedCentralGoogle Scholar
  15. Bretscher MS (2008) On the shape of migrating cells–a ‘front-to-back’ model. J Cell Sci 121:2625–2628PubMedCrossRefGoogle Scholar
  16. Bretscher MS, Aguado-Velasco C (1998) Membrane traffic during cell locomotion. Curr Opin Cell Biol 10:537–541PubMedCrossRefGoogle Scholar
  17. Breuss M, Heng JI, Poirier K, Tian G, Jaglin XH, Qu Z, Braun A, Gstrein T, Ngo L, Haas M, Bahi-Buisson N, Moutard ML, Passemard S, Verloes A, Gressens P, Xie Y, Robson KJ, Rani DS, Thangaraj K, Clausen T, Chelly J, Cowan NJ, Keays DA (2012) Mutations in the beta-tubulin gene TUBB5 cause microcephaly with structural brain abnormalities. Cell Rep 2:1554–1562PubMedPubMedCentralCrossRefGoogle Scholar
  18. Briggs MW, Li Z, Sacks DB (2002) IQGAP1-mediated stimulation of transcriptional co-activation by beta-catenin is modulated by calmodulin. J Biol Chem 277:7453–7465PubMedCrossRefGoogle Scholar
  19. Briggs MW, Sacks DB (2003a) IQGAP1 as signal integrator: Ca2+, calmodulin, Cdc42 and the cytoskeleton. FEBS Lett 542:7–11PubMedCrossRefGoogle Scholar
  20. Briggs MW, Sacks DB (2003b) IQGAP proteins are integral components of cytoskeletal regulation. EMBO Rep 4:571–574PubMedPubMedCentralCrossRefGoogle Scholar
  21. Buck KB, Zheng JQ (2002) Growth cone turning induced by direct local modification of microtubule dynamics. J Neurosci Off J Soc Neurosci 22:9358–9367Google Scholar
  22. Bulinski JC, Gundersen GG (1991) Stabilization of post-translational modification of microtubules during cellular morphogenesis. BioEssays News Rev Mol Cell Develop Biol 13:285–293CrossRefGoogle Scholar
  23. Burack MA, Silverman MA, Banker G (2000) The role of selective transport in neuronal protein sorting. Neuron 26:465–472PubMedCrossRefGoogle Scholar
  24. Cai D, McEwen DP, Martens JR, Meyhofer E, Verhey KJ (2009) Single molecule imaging reveals differences in microtubule track selection between Kinesin motors. PLoS Biol 7:e1000216PubMedPubMedCentralCrossRefGoogle Scholar
  25. Caspi M, Atlas R, Kantor A, Sapir T, Reiner O (2000) Interaction between LIS1 and doublecortin, two lissencephaly gene products. Hum Mol Genet 9:2205–2213PubMedCrossRefGoogle Scholar
  26. Caswell P, Norman J (2008) Endocytic transport of integrins during cell migration and invasion. Trends Cell Biol 18:257–263PubMedCrossRefGoogle Scholar
  27. Cavallaro U, Niedermeyer J, Fuxa M, Christofori G (2001) N-CAM modulates tumour-cell adhesion to matrix by inducing FGF-receptor signalling. Nat Cell Biol 3:650–657PubMedCrossRefGoogle Scholar
  28. Chabin-Brion K, Marceiller J, Perez F, Settegrana C, Drechou A, Durand G, Pous C (2001) The Golgi complex is a microtubule-organizing organelle. Mol Biol Cell 12:2047–2060PubMedPubMedCentralCrossRefGoogle Scholar
  29. Chang YC, Nalbant P, Birkenfeld J, Chang ZF, Bokoch GM (2008) GEF-H1 couples nocodazole-induced microtubule disassembly to cell contractility via RhoA. Mol Biol Cell 19:2147–2153PubMedPubMedCentralCrossRefGoogle Scholar
  30. Chao WT, Kunz J (2009) Focal adhesion disassembly requires clathrin-dependent endocytosis of integrins. FEBS Lett 583:1337–1343PubMedPubMedCentralCrossRefGoogle Scholar
  31. Chen X, S-i K, Borisy GG, Green KJ (2003) p120 catenin associates with kinesin and facilitates the transport of cadherin-catenin complexes to intercellular junctions. J Cell Biol 163:547–557PubMedPubMedCentralCrossRefGoogle Scholar
  32. Chi Z, Melendez AJ (2007) Role of cell adhesion molecules and immune-cell migration in the initiation, onset and development of atherosclerosis. Cell Adh Migr 1:171–175PubMedPubMedCentralCrossRefGoogle Scholar
  33. Chia EW, Grainger R, Harper JL (2008) Colchicine suppresses neutrophil superoxide production in a murine model of gouty arthritis: a rationale for use of low-dose colchicine. Br J Pharmacol 153:1288–1295PubMedPubMedCentralCrossRefGoogle Scholar
  34. Clevers H (2006) Wnt/beta-catenin signaling in development and disease. Cell 127:469–480PubMedCrossRefGoogle Scholar
  35. Colvin RA, Means TK, Diefenbach TJ, Moita LF, Friday RP, Sever S, Campanella GS, Abrazinski T, Manice LA, Moita C, Andrews NW, Wu D, Hacohen N, Luster AD (2010) Synaptotagmin-mediated vesicle fusion regulates cell migration. Nat Immunol 11:495–502PubMedPubMedCentralCrossRefGoogle Scholar
  36. Cooper JA (2013) Cell biology in neuroscience: mechanisms of cell migration in the nervous system. J Cell Biol 202:725–734PubMedPubMedCentralCrossRefGoogle Scholar
  37. Coopman PJ, Do MT, Thompson EW, Mueller SC (1998) Phagocytosis of cross-linked gelatin matrix by human breast carcinoma cells correlates with their invasive capacity. Clin Cancer Res Off J Am Assoc Cancer Res 4:507–515Google Scholar
  38. Cornfine S, Himmel M, Kopp P, El Azzouzi K, Wiesner C, Kruger M, Rudel T, Linder S (2011) The kinesin KIF9 and reggie/flotillin proteins regulate matrix degradation by macrophage podosomes. Mol Biol Cell 22:202–215PubMedPubMedCentralCrossRefGoogle Scholar
  39. Cui DH, Jiang KD, Jiang SD, Xu YF, Yao H (2005) The tumor suppressor adenomatous polyposis coli gene is associated with susceptibility to schizophrenia. Mol Psychiatry 10:669–677PubMedCrossRefGoogle Scholar
  40. de Anda FC, Pollarolo G, Da Silva JS, Camoletto PG, Feiguin F, Dotti CG (2005) Centrosome localization determines neuronal polarity. Nature 436:704–708PubMedCrossRefGoogle Scholar
  41. Del Rio JA, Gonzalez-Billault C, Urena JM, Jimenez EM, Barallobre MJ, Pascual M, Pujadas L, Simo S, La Torre A, Wandosell F, Avila J, Soriano E (2004) MAP1B is required for Netrin 1 signaling in neuronal migration and axonal guidance. Curr Biol CB 14:840–850PubMedCrossRefGoogle Scholar
  42. Dixit R, Ross JL, Goldman YE, Holzbaur EL (2008) Differential regulation of dynein and kinesin motor proteins by tau. Science 319:1086–1089PubMedPubMedCentralCrossRefGoogle Scholar
  43. Drabek K, van Ham M, Stepanova T, Draegestein K, van Horssen R, Sayas CL, Akhmanova A, Ten Hagen T, Smits R, Fodde R, Grosveld F, Galjart N (2006) Role of CLASP2 in microtubule stabilization and the regulation of persistent motility. Curr Biol CB 16:2259–2264PubMedCrossRefGoogle Scholar
  44. Draberova E, Vinopal S, Morfini G, Liu PS, Sladkova V, Sulimenko T, Burns MR, Solowska J, Kulandaivel K, de Chadarevian JP, Legido A, Mork SJ, Janacek J, Baas PW, Draber P, Katsetos CD (2011) Microtubule-severing ATPase spastin in glioblastoma: increased expression in human glioblastoma cell lines and inverse roles in cell motility and proliferation. J Neuropathol Exp Neurol 70:811–826PubMedPubMedCentralCrossRefGoogle Scholar
  45. Duan X, Chang JH, Ge S, Faulkner RL, Kim JY, Kitabatake Y, Liu XB, Yang CH, Jordan JD, Ma DK, Liu CY, Ganesan S, Cheng HJ, Ming GL, Lu B, Song H (2007) Disrupted-In-Schizophrenia 1 regulates integration of newly generated neurons in the adult brain. Cell 130:1146–1158PubMedPubMedCentralCrossRefGoogle Scholar
  46. Dunn GA, Zicha D, Fraylich PE (1997) Rapid, microtubule-dependent fluctuations of the cell margin. J Cell Sci 110(Pt 24):3091–3098PubMedGoogle Scholar
  47. Dupin I, Camand E, Etienne-Manneville S (2009) Classical cadherins control nucleus and centrosome position and cell polarity. J Cell Biol 185:779–786PubMedPubMedCentralCrossRefGoogle Scholar
  48. Dziezanowski MA, DeStefano MJ, Rabinovitch M (1980) Effect of antitubulins on spontaneous and chemotactic migration of neutrophils under agarose. J Cell Sci 42:379–388PubMedGoogle Scholar
  49. Ebneth A, Drewes G, Mandelkow EM, Mandelkow E (1999) Phosphorylation of MAP2c and MAP4 by MARK kinases leads to the destabilization of microtubules in cells. Cell Motil Cytoskeleton 44:209–224PubMedCrossRefGoogle Scholar
  50. Eden S, Rohatgi R, Podtelejnikov AV, Mann M, Kirschner MW (2002) Mechanism of regulation of WAVE1-induced actin nucleation by Rac1 and Nck. Nature 418:790–793PubMedCrossRefGoogle Scholar
  51. Efimov A, Kharitonov A, Efimova N, Loncarek J, Miller PM, Andreyeva N, Gleeson P, Galjart N, Maia AR, McLeod IX, Yates JR 3rd, Maiato H, Khodjakov A, Akhmanova A, Kaverina I (2007) Asymmetric CLASP-dependent nucleation of noncentrosomal microtubules at the trans-Golgi network. Dev Cell 12:917–930PubMedPubMedCentralCrossRefGoogle Scholar
  52. Efimov A, Schiefermeier N, Grigoriev I, Ohi R, Brown MC, Turner CE, Small JV, Kaverina I (2008) Paxillin-dependent stimulation of microtubule catastrophes at focal adhesion sites. J Cell Sci 121:196–204PubMedPubMedCentralCrossRefGoogle Scholar
  53. Efimova N, Grimaldi A, Bachmann A, Frye K, Zhu X, Feoktistov A, Straube A, Kaverina I (2014) Podosome-regulating kinesin KIF1C translocates to the cell periphery in a CLASP-dependent manner. J Cell Sci 127:5179–5188PubMedPubMedCentralCrossRefGoogle Scholar
  54. Errico A, Ballabio A, Rugarli EI (2002) Spastin, the protein mutated in autosomal dominant hereditary spastic paraplegia, is involved in microtubule dynamics. Hum Mol Genet 11:153–163PubMedCrossRefGoogle Scholar
  55. Eskova A, Knapp B, Matelska D, Reusing S, Arjonen A, Lisauskas T, Pepperkok R, Russell R, Eils R, Ivaska J, Kaderali L, Erfle H, Starkuviene V (2014) An RNAi screen identifies KIF15 as a novel regulator of the endocytic trafficking of integrin. J Cell Sci 127:2433–2447PubMedCrossRefGoogle Scholar
  56. Etienne-Manneville S (2004) Actin and microtubules in cell motility: which one is in control? Traffic 5:470–477PubMedCrossRefGoogle Scholar
  57. Etienne-Manneville S (2013) Microtubules in cell migration. Annu Rev Cell Dev Biol 29:471–499PubMedCrossRefGoogle Scholar
  58. Etienne-Manneville S, Hall A (2001) Integrin-mediated activation of Cdc42 controls cell polarity in migrating astrocytes through PKCzeta. Cell 106:489–498PubMedCrossRefGoogle Scholar
  59. Etienne-Manneville S, Hall A (2003) Cdc42 regulates GSK-3beta and adenomatous polyposis coli to control cell polarity. Nature 421:753–756PubMedCrossRefGoogle Scholar
  60. Etienne-Manneville S, Manneville JB, Nicholls S, Ferenczi MA, Hall A (2005) Cdc42 and Par6-PKCzeta regulate the spatially localized association of Dlg1 and APC to control cell polarization. J Cell Biol 170:895–901PubMedPubMedCentralCrossRefGoogle Scholar
  61. Euteneuer U, Schliwa M (1984) Persistent, directional motility of cells and cytoplasmic fragments in the absence of microtubules. Nature 310:58–61PubMedCrossRefGoogle Scholar
  62. Ezratty EJ, Bertaux C, Marcantonio EE, Gundersen GG (2009) Clathrin mediates integrin endocytosis for focal adhesion disassembly in migrating cells. J Cell Biol 187:733–747PubMedPubMedCentralCrossRefGoogle Scholar
  63. Ezratty EJ, Partridge MA, Gundersen GG (2005) Microtubule-induced focal adhesion disassembly is mediated by dynamin and focal adhesion kinase. Nat Cell Biol 7:581–590PubMedCrossRefGoogle Scholar
  64. Fahrion JK, Komuro Y, Li Y, Ohno N, Littner Y, Raoult E, Galas L, Vaudry D, Komuro H (2012) Rescue of neuronal migration deficits in a mouse model of fetal Minamata disease by increasing neuronal Ca2+ spike frequency. Proc Natl Acad Sci U S A 109:5057–5062PubMedPubMedCentralCrossRefGoogle Scholar
  65. Faux MC, Ross JL, Meeker C, Johns T, Ji H, Simpson RJ, Layton MJ, Burgess AW (2004) Restoration of full-length adenomatous polyposis coli (APC) protein in a colon cancer cell line enhances cell adhesion. J Cell Sci 117:427–439PubMedCrossRefGoogle Scholar
  66. Francavilla C, Loeffler S, Piccini D, Kren A, Christofori G, Cavallaro U (2007) Neural cell adhesion molecule regulates the cellular response to fibroblast growth factor. J Cell Sci 120:4388–4394PubMedCrossRefGoogle Scholar
  67. Friedl P, Entschladen F, Conrad C, Niggemann B, Zanker KS (1998a) CD4+ T lymphocytes migrating in three-dimensional collagen lattices lack focal adhesions and utilize beta1 integrin-independent strategies for polarization, interaction with collagen fibers and locomotion. Eur J Immunol 28:2331–2343PubMedCrossRefGoogle Scholar
  68. Friedl P, Gilmour D (2009) Collective cell migration in morphogenesis, regeneration and cancer. Nat Rev Mol Cell Biol 10:445–457PubMedCrossRefGoogle Scholar
  69. Friedl P, Hegerfeldt Y, Tusch M (2004) Collective cell migration in morphogenesis and cancer. Int J Dev Biol 48:441–449PubMedCrossRefGoogle Scholar
  70. Friedl P, Weigelin B (2008) Interstitial leukocyte migration and immune function. Nat Immunol 9:960–969PubMedCrossRefGoogle Scholar
  71. Friedl P, Wolf K (2003) Tumour-cell invasion and migration: diversity and escape mechanisms. Nat Rev Cancer 3:362–374PubMedCrossRefGoogle Scholar
  72. Friedl P, Zanker KS, Brocker EB (1998b) Cell migration strategies in 3-D extracellular matrix: differences in morphology, cell matrix interactions, and integrin function. Microsc Res Tech 43:369–378PubMedCrossRefGoogle Scholar
  73. Fukata M, Kuroda S, Nakagawa M, Kawajiri A, Itoh N, Shoji I, Matsuura Y, Yonehara S, Fujisawa H, Kikuchi A, Kaibuchi K (1999) Cdc42 and Rac1 regulate the interaction of IQGAP1 with beta-catenin. J Biol Chem 274:26044–26050PubMedCrossRefGoogle Scholar
  74. Fukata M, Watanabe T, Noritake J, Nakagawa M, Yamaga M, Kuroda S, Matsuura Y, Iwamatsu A, Perez F, Kaibuchi K (2002) Rac1 and Cdc42 capture microtubules through IQGAP1 and CLIP-170. Cell 109:873–885PubMedCrossRefGoogle Scholar
  75. Ganguly A, Yang H, Sharma R, Patel KD, Cabral F (2012) The role of microtubules and their dynamics in cell migration. J Biol Chem 287:43359–43369PubMedPubMedCentralCrossRefGoogle Scholar
  76. Ghosh-Roy A, Goncharov A, Jin Y, Chisholm AD (2012) Kinesin-13 and tubulin posttranslational modifications regulate microtubule growth in axon regeneration. Dev Cell 23:716–728PubMedPubMedCentralCrossRefGoogle Scholar
  77. Glaven JA, Whitehead I, Bagrodia S, Kay R, Cerione RA (1999) The Dbl-related protein, Lfc, localizes to microtubules and mediates the activation of Rac signaling pathways in cells. J Biol Chem 274:2279–2285PubMedCrossRefGoogle Scholar
  78. Gleeson JG, Lin PT, Flanagan LA, Walsh CA (1999a) Doublecortin is a microtubule-associated protein and is expressed widely by migrating neurons. Neuron 23:257–271PubMedCrossRefGoogle Scholar
  79. Gleeson JG, Minnerath SR, Fox JW, Allen KM, Luo RF, Hong SE, Berg MJ, Kuzniecky R, Reitnauer PJ, Borgatti R, Mira AP, Guerrini R, Holmes GL, Rooney CM, Berkovic S, Scheffer I, Cooper EC, Ricci S, Cusmai R, Crawford TO, Leroy R, Andermann E, Wheless JW, Dobyns WB, Walsh CA et al (1999b) Characterization of mutations in the gene doublecortin in patients with double cortex syndrome. Ann Neurol 45:146–153PubMedCrossRefGoogle Scholar
  80. Godinho SA, Picone R, Burute M, Dagher R, Su Y, Leung CT, Polyak K, Brugge JS, Thery M, Pellman D (2014) Oncogene-like induction of cellular invasion from centrosome amplification. Nature 510:167–171PubMedPubMedCentralCrossRefGoogle Scholar
  81. Gomes ER, Jani S, Gundersen GG (2005) Nuclear movement regulated by Cdc42, MRCK, myosin, and actin flow establishes MTOC polarization in migrating cells. Cell 121:451–463PubMedCrossRefGoogle Scholar
  82. Gu Z, Noss EH, Hsu VW, Brenner MB (2011) Integrins traffic rapidly via circular dorsal ruffles and macropinocytosis during stimulated cell migration. J Cell Biol 193:61–70PubMedPubMedCentralCrossRefGoogle Scholar
  83. Gundersen GG, Bulinski JC (1988) Selective stabilization of microtubules oriented toward the direction of cell migration. Proc Natl Acad Sci U S A 85:5946–5950PubMedPubMedCentralCrossRefGoogle Scholar
  84. Hall A (2012) Rho family GTPases. Biochem Soc Trans 40:1378–1382PubMedCrossRefGoogle Scholar
  85. Hamshere ML, Walters JT, Smith R, Richards AL, Green E, Grozeva D, Jones I, Forty L, Jones L, Gordon-Smith K, Riley B, O’Neill FA, Kendler KS, Sklar P, Purcell S, Kranz J, Schizophrenia Psychiatric Genome-wide Association Study C, Wellcome Trust Case Control C, Wellcome Trust Case Control C, Morris D, Gill M, Holmans P, Craddock N, Corvin A, Owen MJ, O’Donovan MC (2013) Genome-wide significant associations in schizophrenia to ITIH3/4, CACNA1C and SDCCAG8, and extensive replication of associations reported by the Schizophrenia PGC. Mol Psychiatry 18:708–712Google Scholar
  86. Harada T, Swift J, Irianto J, Shin JW, Spinler KR, Athirasala A, Diegmiller R, Dingal PC, Ivanovska IL, Discher DE (2014) Nuclear lamin stiffness is a barrier to 3D migration, but softness can limit survival. J Cell Biol 204:669–682PubMedPubMedCentralCrossRefGoogle Scholar
  87. Hashimoto R, Numakawa T, Ohnishi T, Kumamaru E, Yagasaki Y, Ishimoto T, Mori T, Nemoto K, Adachi N, Izumi A, Chiba S, Noguchi H, Suzuki T, Iwata N, Ozaki N, Taguchi T, Kamiya A, Kosuga A, Tatsumi M, Kamijima K, Weinberger DR, Sawa A, Kunugi H (2006) Impact of the DISC1 Ser704Cys polymorphism on risk for major depression, brain morphology and ERK signaling. Hum Mol Genet 15:3024–3033PubMedCrossRefGoogle Scholar
  88. Hattori M, Adachi H, Tsujimoto M, Arai H, Inoue K (1994) Miller-Dieker lissencephaly gene encodes a subunit of brain platelet-activating factor acetylhydrolase [corrected]. Nature 370:216–218PubMedCrossRefGoogle Scholar
  89. Hayashi K, Suzuki A, Ohno S (2012) PAR-1/MARK: a kinase essential for maintaining the dynamic state of microtubules. Cell Struct Funct 37:21–25PubMedCrossRefGoogle Scholar
  90. Hegerfeldt Y, Tusch M, Brocker EB, Friedl P (2002) Collective cell movement in primary melanoma explants: plasticity of cell-cell interaction, beta1-integrin function, and migration strategies. Cancer Res 62:2125–2130PubMedGoogle Scholar
  91. Heintz TG, Heller J, Zhao R, Caceres A, Eva R, Fawcett JW (2014) Kinesin KIF4A transports integrin beta1 in developing axons of cortical neurons. Mol Cell Neurosci 63:60–71PubMedCrossRefGoogle Scholar
  92. Hennah W, Thomson P, McQuillin A, Bass N, Loukola A, Anjorin A, Blackwood D, Curtis D, Deary IJ, Harris SE, Isometsa ET, Lawrence J, Lonnqvist J, Muir W, Palotie A, Partonen T, Paunio T, Pylkko E, Robinson M, Soronen P, Suominen K, Suvisaari J, Thirumalai S, St Clair D, Gurling H, Peltonen L, Porteous D (2009) DISC1 association, heterogeneity and interplay in schizophrenia and bipolar disorder. Mol Psychiatry 14:865–873PubMedCrossRefGoogle Scholar
  93. Heuberger J, Birchmeier W (2010) Interplay of cadherin-mediated cell adhesion and canonical Wnt signaling. Cold Spring Harb Perspect Biol 2:a002915PubMedPubMedCentralCrossRefGoogle Scholar
  94. Huang CF, Banker G (2012) The translocation selectivity of the kinesins that mediate neuronal organelle transport. Traffic 13:549–564PubMedPubMedCentralCrossRefGoogle Scholar
  95. Humbert PO, Grzeschik NA, Brumby AM, Galea R, Elsum I, Richardson HE (2008) Control of tumourigenesis by the Scribble/Dlg/Lgl polarity module. Oncogene 27:6888–6907PubMedCrossRefGoogle Scholar
  96. Hynes RO (2002) Integrins: bidirectional, allosteric signaling machines. Cell 110:673–687PubMedCrossRefGoogle Scholar
  97. Ichii T, Takeichi M (2007) p120-catenin regulates microtubule dynamics and cell migration in a cadherin-independent manner. Genes Cells Devoted Mol Cell Mech 12:827–839CrossRefGoogle Scholar
  98. Insolera R, Shao W, Airik R, Hildebrandt F, Shi SH (2014) SDCCAG8 regulates pericentriolar material recruitment and neuronal migration in the developing cortex. Neuron 83:805–822PubMedPubMedCentralCrossRefGoogle Scholar
  99. Ishizuka K, Kamiya A, Oh EC, Kanki H, Seshadri S, Robinson JF, Murdoch H, Dunlop AJ, Kubo K, Furukori K, Huang B, Zeledon M, Hayashi-Takagi A, Okano H, Nakajima K, Houslay MD, Katsanis N, Sawa A (2011) DISC1-dependent switch from progenitor proliferation to migration in the developing cortex. Nature 473:92–96PubMedPubMedCentralCrossRefGoogle Scholar
  100. Jacobson C, Schnapp B, Banker GA (2006) A change in the selective translocation of the Kinesin-1 motor domain marks the initial specification of the axon. Neuron 49:797–804PubMedCrossRefGoogle Scholar
  101. Jaglin XH, Poirier K, Saillour Y, Buhler E, Tian G, Bahi-Buisson N, Fallet-Bianco C, Phan-Dinh-Tuy F, Kong XP, Bomont P, Castelnau-Ptakhine L, Odent S, Loget P, Kossorotoff M, Snoeck I, Plessis G, Parent P, Beldjord C, Cardoso C, Represa A, Flint J, Keays DA, Cowan NJ, Chelly J (2009) Mutations in the beta-tubulin gene TUBB2B result in asymmetrical polymicrogyria. Nat Genet 41:746–752PubMedPubMedCentralCrossRefGoogle Scholar
  102. Jaulin F, Kreitzer G (2010) KIF17 stabilizes microtubules and contributes to epithelial morphogenesis by acting at MT plus ends with EB1 and APC. J Cell Biol 190:443–460PubMedPubMedCentralCrossRefGoogle Scholar
  103. Jenkins B, Decker H, Bentley M, Luisi J, Banker G (2012) A novel split kinesin assay identifies motor proteins that interact with distinct vesicle populations. J Cell Biol 198:749–761PubMedPubMedCentralCrossRefGoogle Scholar
  104. Jiang K, Hua S, Mohan R, Grigoriev I, Yau KW, Liu Q, Katrukha EA, Altelaar AF, Heck AJ, Hoogenraad CC, Akhmanova A (2014) Microtubule minus-end stabilization by polymerization-driven CAMSAP deposition. Dev Cell 28:295–309PubMedCrossRefGoogle Scholar
  105. Jolly AL, Kim H, Srinivasan D, Lakonishok M, Larson AG, Gelfand VI (2010) Kinesin-1 heavy chain mediates microtubule sliding to drive changes in cell shape. Proc Natl Acad Sci U S A 107:12151–12156PubMedPubMedCentralCrossRefGoogle Scholar
  106. Jordan MA, Wilson L (2004) Microtubules as a target for anticancer drugs. Nat Rev Cancer 4:253–265PubMedCrossRefGoogle Scholar
  107. Jossin Y, Cooper JA (2011) Reelin, Rap1 and N-cadherin orient the migration of multipolar neurons in the developing neocortex. Nat Neurosci 14:697–703PubMedPubMedCentralCrossRefGoogle Scholar
  108. Kamiya A, Tan PL, Kubo K, Engelhard C, Ishizuka K, Kubo A, Tsukita S, Pulver AE, Nakajima K, Cascella NG, Katsanis N, Sawa A (2008) Recruitment of PCM1 to the centrosome by the cooperative action of DISC1 and BBS4: a candidate for psychiatric illnesses. Arch Gen Psychiatry 65:996–1006PubMedPubMedCentralCrossRefGoogle Scholar
  109. Kassler S, Donninger H, Birrer MJ, Clark GJ (2012) RASSF1A and the taxol response in ovarian cancer. Mol Biol Int 2012:263267PubMedPubMedCentralCrossRefGoogle Scholar
  110. Katz ZB, Wells AL, Park HY, Wu B, Shenoy SM, Singer RH (2012) beta-Actin mRNA compartmentalization enhances focal adhesion stability and directs cell migration. Genes Dev 26:1885–1890PubMedPubMedCentralCrossRefGoogle Scholar
  111. Kaverina I, Krylyshkina O, Small JV (1999) Microtubule targeting of substrate contacts promotes their relaxation and dissociation. J Cell Biol 146:1033–1044PubMedPubMedCentralCrossRefGoogle Scholar
  112. Kaverina I, Rottner K, Small JV (1998) Targeting, capture, and stabilization of microtubules at early focal adhesions. J Cell Biol 142:181–190PubMedPubMedCentralCrossRefGoogle Scholar
  113. Kaverina I, Straube A (2011) Regulation of cell migration by dynamic microtubules. Semin Cell Dev Biol 22:968–974PubMedPubMedCentralCrossRefGoogle Scholar
  114. Kawauchi T, Sekine K, Shikanai M, Chihama K, Tomita K, Kubo K, Nakajima K, Nabeshima Y, Hoshino M (2010) Rab GTPases-dependent endocytic pathways regulate neuronal migration and maturation through N-cadherin trafficking. Neuron 67:588–602PubMedCrossRefGoogle Scholar
  115. Keays DA, Tian G, Poirier K, Huang GJ, Siebold C, Cleak J, Oliver PL, Fray M, Harvey RJ, Molnar Z, Pinon MC, Dear N, Valdar W, Brown SD, Davies KE, Rawlins JN, Cowan NJ, Nolan P, Chelly J, Flint J (2007) Mutations in alpha-tubulin cause abnormal neuronal migration in mice and lissencephaly in humans. Cell 128:45–57PubMedPubMedCentralCrossRefGoogle Scholar
  116. Keren K, Pincus Z, Allen GM, Barnhart EL, Marriott G, Mogilner A, Theriot JA (2008) Mechanism of shape determination in motile cells. Nature 453:475–480PubMedPubMedCentralCrossRefGoogle Scholar
  117. Kholmanskikh SS, Koeller HB, Wynshaw-Boris A, Gomez T, Letourneau PC, Ross ME (2006) Calcium-dependent interaction of Lis1 with IQGAP1 and Cdc42 promotes neuronal motility. Nat Neurosci 9:50–57PubMedCrossRefGoogle Scholar
  118. Kislauskis EH, Zhu X, Singer RH (1997) beta-Actin messenger RNA localization and protein synthesis augment cell motility. J Cell Biol 136:1263–1270PubMedPubMedCentralCrossRefGoogle Scholar
  119. Kita K, Wittmann T, Nathke IS, Waterman-Storer CM (2006) Adenomatous polyposis coli on microtubule plus ends in cell extensions can promote microtubule net growth with or without EB1. Mol Biol Cell 17:2331–2345PubMedPubMedCentralCrossRefGoogle Scholar
  120. Kodama A, Karakesisoglou I, Wong E, Vaezi A, Fuchs E (2003) ACF7: an essential integrator of microtubule dynamics. Cell 115:343–354PubMedCrossRefGoogle Scholar
  121. Komarova YA, Vorobjev IA, Borisy GG (2002) Life cycle of MTs: persistent growth in the cell interior, asymmetric transition frequencies and effects of the cell boundary. J Cell Sci 115:3527–3539PubMedGoogle Scholar
  122. Komuro H, Rakic P (1998) Distinct modes of neuronal migration in different domains of developing cerebellar cortex. J Neurosci Off J Soc Neurosci 18:1478–1490Google Scholar
  123. Kopp P, Lammers R, Aepfelbacher M, Woehlke G, Rudel T, Machuy N, Steffen W, Linder S (2006) The kinesin KIF1C and microtubule plus ends regulate podosome dynamics in macrophages. Mol Biol Cell 17:2811–2823PubMedPubMedCentralCrossRefGoogle Scholar
  124. Kozlovsky N, Belmaker RH, Agam G (2002) GSK-3 and the neurodevelopmental hypothesis of schizophrenia. Eur Neuropsychopharmacol J Eur Coll Neuropsychopharmacol 12:13–25CrossRefGoogle Scholar
  125. Krendel M, Zenke FT, Bokoch GM (2002) Nucleotide exchange factor GEF-H1 mediates cross-talk between microtubules and the actin cytoskeleton. Nat Cell Biol 4:294–301PubMedCrossRefGoogle Scholar
  126. Kroboth K, Newton IP, Kita K, Dikovskaya D, Zumbrunn J, Waterman-Storer CM, Nathke IS (2007) Lack of adenomatous polyposis coli protein correlates with a decrease in cell migration and overall changes in microtubule stability. Mol Biol Cell 18:910–918PubMedPubMedCentralCrossRefGoogle Scholar
  127. Krylyshkina O, Anderson KI, Kaverina I, Upmann I, Manstein DJ, Small JV, Toomre DK (2003) Nanometer targeting of microtubules to focal adhesions. J Cell Biol 161:853–859PubMedPubMedCentralCrossRefGoogle Scholar
  128. Krylyshkina O, Kaverina I, Kranewitter W, Steffen W, Alonso MC, Cross RA, Small JV (2002) Modulation of substrate adhesion dynamics via microtubule targeting requires kinesin-1. J Cell Biol 156:349–359PubMedPubMedCentralCrossRefGoogle Scholar
  129. Kumar P, Lyle KS, Gierke S, Matov A, Danuser G, Wittmann T (2009) GSK3beta phosphorylation modulates CLASP-microtubule association and lamella microtubule attachment. J Cell Biol 184:895–908PubMedPubMedCentralCrossRefGoogle Scholar
  130. Kumar S, Xu J, Perkins C, Guo F, Snapper S, Finkelman FD, Zheng Y, Filippi MD (2012) Cdc42 regulates neutrophil migration via crosstalk between WASp, CD11b, and microtubules. Blood 120:3563–3574PubMedPubMedCentralCrossRefGoogle Scholar
  131. Kupfer A, Dennert G, Singer SJ (1983) Polarization of the Golgi apparatus and the microtubule-organizing center within cloned natural killer cells bound to their targets. Proc Natl Acad Sci U S A 80:7224–7228PubMedPubMedCentralCrossRefGoogle Scholar
  132. Kushner EJ, Ferro LS, Liu JY, Durrant JR, Rogers SL, Dudley AC, Bautch VL (2014) Excess centrosomes disrupt endothelial cell migration via centrosome scattering. J Cell Biol 206:257–272PubMedPubMedCentralCrossRefGoogle Scholar
  133. Lacroix B, van Dijk J, Gold ND, Guizetti J, Aldrian-Herrada G, Rogowski K, Gerlich DW, Janke C (2010) Tubulin polyglutamylation stimulates spastin-mediated microtubule severing. J Cell Biol 189:945–954PubMedPubMedCentralCrossRefGoogle Scholar
  134. Lammermann T, Sixt M (2009) Mechanical modes of ‘amoeboid’ cell migration. Curr Opin Cell Biol 21:636–644PubMedCrossRefGoogle Scholar
  135. Lawrence JB, Singer RH (1986) Intracellular localization of messenger RNAs for cytoskeletal proteins. Cell 45:407–415PubMedCrossRefGoogle Scholar
  136. Lee G, Leugers CJ (2012) Tau and tauopathies. Prog Mol Biol Transl Sci 107:263–293PubMedPubMedCentralCrossRefGoogle Scholar
  137. Legate KR, Montanez E, Kudlacek O, Fassler R (2006) ILK, PINCH and parvin: the tIPP of integrin signalling. Nat Rev Mol Cell Biol 7:20–31PubMedCrossRefGoogle Scholar
  138. Lehembre F, Yilmaz M, Wicki A, Schomber T, Strittmatter K, Ziegler D, Kren A, Went P, Derksen PW, Berns A, Jonkers J, Christofori G (2008) NCAM-induced focal adhesion assembly: a functional switch upon loss of E-cadherin. EMBO J 27:2603–2615PubMedPubMedCentralCrossRefGoogle Scholar
  139. Lele Z, Folchert A, Concha M, Rauch GJ, Geisler R, Rosa F, Wilson SW, Hammerschmidt M, Bally-Cuif L (2002) parachute/n-cadherin is required for morphogenesis and maintained integrity of the zebrafish neural tube. Development 129:3281–3294PubMedGoogle Scholar
  140. Li D, Xie S, Ren Y, Huo L, Gao J, Cui D, Liu M, Zhou J (2011) Microtubule-associated deacetylase HDAC6 promotes angiogenesis by regulating cell migration in an EB1-dependent manner. Protein Cell 2:150–160PubMedCrossRefGoogle Scholar
  141. Liao G, Nagasaki T, Gundersen GG (1995) Low concentrations of nocodazole interfere with fibroblast locomotion without significantly affecting microtubule level: implications for the role of dynamic microtubules in cell locomotion. J Cell Sci 108(Pt 11):3473–3483PubMedGoogle Scholar
  142. Lindeboom JJ, Nakamura M, Hibbel A, Shundyak K, Gutierrez R, Ketelaar T, Emons AM, Mulder BM, Kirik V, Ehrhardt DW (2013) A mechanism for reorientation of cortical microtubule arrays driven by microtubule severing. Science 342:1245533PubMedCrossRefGoogle Scholar
  143. Liu JS (2011) Molecular genetics of neuronal migration disorders. Curr Neurol Neurosci Rep 11:171–178PubMedCrossRefGoogle Scholar
  144. Liu M, Nadar VC, Kozielski F, Kozlowska M, Yu W, Baas PW (2010) Kinesin-12, a mitotic microtubule-associated motor protein, impacts axonal growth, navigation, and branching. J Neurosci Off J Soc Neurosci 30:14896–14906CrossRefGoogle Scholar
  145. Liu R, Woolner S, Johndrow JE, Metzger D, Flores A, Parkhurst SM (2008) Sisyphus, the Drosophila myosin XV homolog, traffics within filopodia transporting key sensory and adhesion cargos. Development 135:53–63PubMedCrossRefGoogle Scholar
  146. Lu W, Fox P, Lakonishok M, Davidson MW, Gelfand VI (2013) Initial neurite outgrowth in Drosophila neurons is driven by kinesin-powered microtubule sliding. Curr Biol CB 23:1018–1023PubMedCrossRefGoogle Scholar
  147. Luxton GW, Gundersen GG (2011) Orientation and function of the nuclear-centrosomal axis during cell migration. Curr Opin Cell Biol 23:579–588PubMedPubMedCentralCrossRefGoogle Scholar
  148. Ma B, Savas JN, Yu MS, Culver BP, Chao MV, Tanese N (2011) Huntingtin mediates dendritic transport of beta-actin mRNA in rat neurons. Sci Rep 1:140PubMedPubMedCentralCrossRefGoogle Scholar
  149. Machacek M, Hodgson L, Welch C, Elliott H, Pertz O, Nalbant P, Abell A, Johnson GL, Hahn KM, Danuser G (2009) Coordination of Rho GTPase activities during cell protrusion. Nature 461:99–103PubMedPubMedCentralCrossRefGoogle Scholar
  150. Mandeville JT, Lawson MA, Maxfield FR (1997) Dynamic imaging of neutrophil migration in three dimensions: mechanical interactions between cells and matrix. J Leukoc Biol 61:188–200PubMedGoogle Scholar
  151. Maretzky T, Reiss K, Ludwig A, Buchholz J, Scholz F, Proksch E, de Strooper B, Hartmann D, Saftig P (2005) ADAM10 mediates E-cadherin shedding and regulates epithelial cell-cell adhesion, migration, and beta-catenin translocation. Proc Natl Acad Sci U S A 102:9182–9187PubMedPubMedCentralCrossRefGoogle Scholar
  152. Margadant C, Monsuur HN, Norman JC, Sonnenberg A (2011) Mechanisms of integrin activation and trafficking. Curr Opin Cell Biol 23:607–614PubMedCrossRefGoogle Scholar
  153. Mary S, Charrasse S, Meriane M, Comunale F, Travo P, Blangy A, Gauthier-Rouviere C (2002) Biogenesis of N-cadherin-dependent cell-cell contacts in living fibroblasts is a microtubule-dependent kinesin-driven mechanism. Mol Biol Cell 13:285–301PubMedPubMedCentralCrossRefGoogle Scholar
  154. Matsumoto S, Fumoto K, Okamoto T, Kaibuchi K, Kikuchi A (2010) Binding of APC and dishevelled mediates Wnt5a-regulated focal adhesion dynamics in migrating cells. EMBO J 29:1192–1204PubMedPubMedCentralCrossRefGoogle Scholar
  155. Matsushima K, Tokuraku K, Hasan MR, Kotani S (2012) Microtubule-associated protein 4 binds to actin filaments and modulates their properties. J Biochem 151:99–108PubMedCrossRefGoogle Scholar
  156. McCrea PD, Gu D, Balda MS (2009) Junctional music that the nucleus hears: cell-cell contact signaling and the modulation of gene activity. Cold Spring Harb Perspect Biol 1:a002923PubMedPubMedCentralCrossRefGoogle Scholar
  157. McLean GW, Carragher NO, Avizienyte E, Evans J, Brunton VG, Frame MC (2005) The role of focal-adhesion kinase in cancer - a new therapeutic opportunity. Nat Rev Cancer 5:505–515PubMedCrossRefGoogle Scholar
  158. Mehlen P, Puisieux A (2006) Metastasis: a question of life or death. Nat Rev Cancer 6:449–458PubMedCrossRefGoogle Scholar
  159. Meyer KD, Morris JA (2009) Disc1 regulates granule cell migration in the developing hippocampus. Hum Mol Genet 18:3286–3297PubMedPubMedCentralCrossRefGoogle Scholar
  160. Mierke CT, Kollmannsberger P, Zitterbart DP, Diez G, Koch TM, Marg S, Ziegler WH, Goldmann WH, Fabry B (2010) Vinculin facilitates cell invasion into three-dimensional collagen matrices. J Biol Chem 285:13121–13130PubMedPubMedCentralCrossRefGoogle Scholar
  161. Miller AL, Wang Y, Mooseker MS, Koleske AJ (2004) The Abl-related gene (Arg) requires its F-actin-microtubule cross-linking activity to regulate lamellipodial dynamics during fibroblast adhesion. J Cell Biol 165:407–419PubMedPubMedCentralCrossRefGoogle Scholar
  162. Miller PM, Folkmann AW, Maia AR, Efimova N, Efimov A, Kaverina I (2009) Golgi-derived CLASP-dependent microtubules control Golgi organization and polarized trafficking in motile cells. Nat Cell Biol 11:1069–1080PubMedPubMedCentralCrossRefGoogle Scholar
  163. Mimori-Kiyosue Y, Shiina N, Tsukita S (2000) Adenomatous polyposis coli (APC) protein moves along microtubules and concentrates at their growing ends in epithelial cells. J Cell Biol 148:505–518PubMedPubMedCentralCrossRefGoogle Scholar
  164. Mingle LA, Okuhama NN, Shi J, Singer RH, Condeelis J, Liu G (2005) Localization of all seven messenger RNAs for the actin-polymerization nucleator Arp2/3 complex in the protrusions of fibroblasts. J Cell Sci 118:2425–2433PubMedPubMedCentralCrossRefGoogle Scholar
  165. Mohn JL, Alexander J, Pirone A, Palka CD, Lee SY, Mebane L, Haydon PG, Jacob MH (2014) Adenomatous polyposis coli protein deletion leads to cognitive and autism-like disabilities. Mol Psychiatry 19:1133–1142PubMedPubMedCentralCrossRefGoogle Scholar
  166. Monier-Gavelle F, Duband JL (1995) Control of N-cadherin-mediated intercellular adhesion in migrating neural crest cells in vitro. J Cell Sci 108(Pt 12):3839–3853PubMedGoogle Scholar
  167. Montenegro-Venegas C, Tortosa E, Rosso S, Peretti D, Bollati F, Bisbal M, Jausoro I, Avila J, Caceres A, Gonzalez-Billault C (2010) MAP1B regulates axonal development by modulating Rho-GTPase Rac1 activity. Mol Biol Cell 21:3518–3528PubMedPubMedCentralCrossRefGoogle Scholar
  168. Munemitsu S, Albert I, Souza B, Rubinfeld B, Polakis P (1995) Regulation of intracellular beta-catenin levels by the adenomatous polyposis coli (APC) tumor-suppressor protein. Proc Natl Acad Sci U S A 92:3046–3050PubMedPubMedCentralCrossRefGoogle Scholar
  169. Myers KA, Baas PW (2007) Kinesin-5 regulates the growth of the axon by acting as a brake on its microtubule array. J Cell Biol 178:1081–1091PubMedPubMedCentralCrossRefGoogle Scholar
  170. Nadar VC, Ketschek A, Myers KA, Gallo G, Baas PW (2008) Kinesin-5 is essential for growth-cone turning. Curr Biol 18(24):1972–1977PubMedPubMedCentralCrossRefGoogle Scholar
  171. Nakagawa S, Takeichi M (1998) Neural crest emigration from the neural tube depends on regulated cadherin expression. Development 125:2963–2971PubMedGoogle Scholar
  172. Nakamura M, Zhou XZ, Lu KP (2001) Critical role for the EB1 and APC interaction in the regulation of microtubule polymerization. Curr Biol CB 11:1062–1067PubMedCrossRefGoogle Scholar
  173. Nalbant P, Chang YC, Birkenfeld J, Chang ZF, Bokoch GM (2009) Guanine nucleotide exchange factor-H1 regulates cell migration via localized activation of RhoA at the leading edge. Mol Biol Cell 20:4070–4082PubMedPubMedCentralCrossRefGoogle Scholar
  174. Nathke IS, Adams CL, Polakis P, Sellin JH, Nelson WJ (1996) The adenomatous polyposis coli tumor suppressor protein localizes to plasma membrane sites involved in active cell migration. J Cell Biol 134:165–179PubMedCrossRefGoogle Scholar
  175. Niethammer P, Bastiaens P, Karsenti E (2004) Stathmin-tubulin interaction gradients in motile and mitotic cells. Science 303:1862–1866PubMedCrossRefGoogle Scholar
  176. Niggli V (2003) Microtubule-disruption-induced and chemotactic-peptide-induced migration of human neutrophils: implications for differential sets of signalling pathways. J Cell Sci 116:813–822PubMedCrossRefGoogle Scholar
  177. Nishimura T, Kaibuchi K (2007) Numb controls integrin endocytosis for directional cell migration with aPKC and PAR-3. Dev Cell 13:15–28PubMedCrossRefGoogle Scholar
  178. Nobes CD, Hall A (1999) Rho GTPases control polarity, protrusion, and adhesion during cell movement. J Cell Biol 144:1235–1244PubMedPubMedCentralCrossRefGoogle Scholar
  179. O’Donnell L, Rhodes D, Smith SJ, Merriner DJ, Clark BJ, Borg C, Whittle B, O’Connor AE, Smith LB, McNally FJ, de Kretser DM, Goodnow CC, Ormandy CJ, Jamsai D, O’Bryan MK (2012) An essential role for katanin p80 and microtubule severing in male gamete production. PLoS Genet 8:e1002698PubMedPubMedCentralCrossRefGoogle Scholar
  180. O’Sullivan D, Miller JH, Northcote PT, La Flamme AC (2013) Microtubule-stabilizing agents delay the onset of EAE through inhibition of migration. Immunol Cell Biol 91:583–592PubMedCrossRefGoogle Scholar
  181. Okada K, Bartolini F, Deaconescu AM, Moseley JB, Dogic Z, Grigorieff N, Gundersen GG, Goode BL (2010) Adenomatous polyposis coli protein nucleates actin assembly and synergizes with the formin mDia1. J Cell Biol 189:1087–1096PubMedPubMedCentralCrossRefGoogle Scholar
  182. Oleynikov Y, Singer RH (1998) RNA localization: different zipcodes, same postman? Trends Cell Biol 8:381–383PubMedCrossRefGoogle Scholar
  183. Osmani N, Peglion F, Chavrier P, Etienne-Manneville S (2010) Cdc42 localization and cell polarity depend on membrane traffic. J Cell Biol 191:1261–1269PubMedPubMedCentralCrossRefGoogle Scholar
  184. Ozmen M, Yilmaz Y, Caliskan M, Minareci O, Aydinli N (2000) Clinical features of 21 patients with lissencephaly type I (agyria-pachygyria). Turk J Pediatr 42:210–214PubMedGoogle Scholar
  185. Palamidessi A, Frittoli E, Garre M, Faretta M, Mione M, Testa I, Diaspro A, Lanzetti L, Scita G, Di Fiore PP (2008) Endocytic trafficking of Rac is required for the spatial restriction of signaling in cell migration. Cell 134:135–147PubMedCrossRefGoogle Scholar
  186. Palazzo AF, Eng CH, Schlaepfer DD, Marcantonio EE, Gundersen GG (2004) Localized stabilization of microtubules by integrin- and FAK-facilitated Rho signaling. Science 303:836–839PubMedCrossRefGoogle Scholar
  187. Palazzo AF, Joseph HL, Chen YJ, Dujardin DL, Alberts AS, Pfister KK, Vallee RB, Gundersen GG (2001) Cdc42, dynein, and dynactin regulate MTOC reorientation independent of Rho-regulated microtubule stabilization. Curr Biol CB 11:1536–1541PubMedCrossRefGoogle Scholar
  188. Paratcha G, Ledda F, Ibanez CF (2003) The neural cell adhesion molecule NCAM is an alternative signaling receptor for GDNF family ligands. Cell 113:867–879PubMedCrossRefGoogle Scholar
  189. Parsons JT, Horwitz AR, Schwartz MA (2010) Cell adhesion: integrating cytoskeletal dynamics and cellular tension. Nat Rev Mol Cell Biol 11:633–643PubMedPubMedCentralCrossRefGoogle Scholar
  190. Pegtel DM, Ellenbroek SI, Mertens AE, van der Kammen RA, de Rooij J, Collard JG (2007) The Par-Tiam1 complex controls persistent migration by stabilizing microtubule-dependent front-rear polarity. Curr Biol CB 17:1623–1634PubMedCrossRefGoogle Scholar
  191. Peris L, Wagenbach M, Lafanechere L, Brocard J, Moore AT, Kozielski F, Job D, Wordeman L, Andrieux A (2009) Motor-dependent microtubule disassembly driven by tubulin tyrosination. J Cell Biol 185:1159–1166PubMedPubMedCentralCrossRefGoogle Scholar
  192. Petrie RJ, Doyle AD, Yamada KM (2009) Random versus directionally persistent cell migration. Nat Rev Mol Cell Biol 10:538–549PubMedPubMedCentralCrossRefGoogle Scholar
  193. Pfister AS, Hadjihannas MV, Rohrig W, Schambony A, Behrens J (2012) Amer2 protein interacts with EB1 protein and adenomatous polyposis coli (APC) and controls microtubule stability and cell migration. J Biol Chem 287:35333–35340PubMedPubMedCentralCrossRefGoogle Scholar
  194. Pilz DT, Matsumoto N, Minnerath S, Mills P, Gleeson JG, Allen KM, Walsh CA, Barkovich AJ, Dobyns WB, Ledbetter DH, Ross ME (1998) LIS1 and XLIS (DCX) mutations cause most classical lissencephaly, but different patterns of malformation. Hum Mol Genet 7:2029–2037PubMedCrossRefGoogle Scholar
  195. Poirier K, Keays DA, Francis F, Saillour Y, Bahi N, Manouvrier S, Fallet-Bianco C, Pasquier L, Toutain A, Tuy FP, Bienvenu T, Joriot S, Odent S, Ville D, Desguerre I, Goldenberg A, Moutard ML, Fryns JP, van Esch H, Harvey RJ, Siebold C, Flint J, Beldjord C, Chelly J (2007) Large spectrum of lissencephaly and pachygyria phenotypes resulting from de novo missense mutations in tubulin alpha 1A (TUBA1A). Hum Mutat 28:1055–1064PubMedCrossRefGoogle Scholar
  196. Poirier K, Lebrun N, Broix L, Tian G, Saillour Y, Boscheron C, Parrini E, Valence S, Pierre BS, Oger M, Lacombe D, Genevieve D, Fontana E, Darra F, Cances C, Barth M, Bonneau D, Bernadina BD, N’Guyen S, Gitiaux C, Parent P, des Portes V, Pedespan JM, Legrez V, Castelnau-Ptakine L, Nitschke P, Hieu T, Masson C, Zelenika D, Andrieux A, Francis F, Guerrini R, Cowan NJ, Bahi-Buisson N, Chelly J (2013) Mutations in TUBG1, DYNC1H1, KIF5C and KIF2A cause malformations of cortical development and microcephaly. Nat Genet 45:639–647PubMedCrossRefGoogle Scholar
  197. Poirier K, Saillour Y, Bahi-Buisson N, Jaglin XH, Fallet-Bianco C, Nabbout R, Castelnau-Ptakhine L, Roubertie A, Attie-Bitach T, Desguerre I, Genevieve D, Barnerias C, Keren B, Lebrun N, Boddaert N, Encha-Razavi F, Chelly J (2010) Mutations in the neuronal ss-tubulin subunit TUBB3 result in malformation of cortical development and neuronal migration defects. Hum Mol Genet 19:4462–4473PubMedPubMedCentralCrossRefGoogle Scholar
  198. Pouthas F, Girard P, Lecaudey V, Ly TB, Gilmour D, Boulin C, Pepperkok R, Reynaud EG (2008) In migrating cells, the Golgi complex and the position of the centrosome depend on geometrical constraints of the substratum. J Cell Sci 121:2406–2414PubMedCrossRefGoogle Scholar
  199. Qi J, Wang J, Romanyuk O, Siu CH (2006) Involvement of Src family kinases in N-cadherin phosphorylation and beta-catenin dissociation during transendothelial migration of melanoma cells. Mol Biol Cell 17:1261–1272PubMedPubMedCentralCrossRefGoogle Scholar
  200. Rappl A, Piontek G, Schlegel J (2008) EGFR-dependent migration of glial cells is mediated by reorganisation of N-cadherin. J Cell Sci 121:4089–4097PubMedCrossRefGoogle Scholar
  201. Ratner S, Sherrod WS, Lichlyter D (1997) Microtubule retraction into the uropod and its role in T cell polarization and motility. J Immunol 159:1063–1067PubMedGoogle Scholar
  202. Recher C, Ysebaert L, Beyne-Rauzy O, Mansat-De Mas V, Ruidavets JB, Cariven P, Demur C, Payrastre B, Laurent G, Racaud-Sultan C (2004) Expression of focal adhesion kinase in acute myeloid leukemia is associated with enhanced blast migration, increased cellularity, and poor prognosis. Cancer Res 64:3191–3197PubMedCrossRefGoogle Scholar
  203. Reed NA, Cai D, Blasius TL, Jih GT, Meyhofer E, Gaertig J, Verhey KJ (2006) Microtubule acetylation promotes kinesin-1 binding and transport. Curr Biol CB 16:2166–2172PubMedCrossRefGoogle Scholar
  204. Ren Y, Li R, Zheng Y, Busch H (1998) Cloning and characterization of GEF-H1, a microtubule-associated guanine nucleotide exchange factor for Rac and Rho GTPases. J Biol Chem 273:34954–34960PubMedCrossRefGoogle Scholar
  205. Revenu C, Streichan S, Dona E, Lecaudey V, Hufnagel L, Gilmour D (2014) Quantitative cell polarity imaging defines leader-to-follower transitions during collective migration and the key role of microtubule-dependent adherens junction formation. Development 141:1282–1291PubMedCrossRefGoogle Scholar
  206. Rid R, Schiefermeier N, Grigoriev I, Small JV, Kaverina I (2005) The last but not the least: the origin and significance of trailing adhesions in fibroblastic cells. Cell Motil Cytoskeleton 61:161–171PubMedCrossRefGoogle Scholar
  207. Ridley AJ, Schwartz MA, Burridge K, Firtel RA, Ginsberg MH, Borisy G, Parsons JT, Horwitz AR (2003) Cell migration: integrating signals from front to back. Science 302:1704–1709PubMedCrossRefGoogle Scholar
  208. Riederer BM (2007) Microtubule-associated protein 1B, a growth-associated and phosphorylated scaffold protein. Brain Res Bull 71:541–558PubMedCrossRefGoogle Scholar
  209. Rieger S, Senghaas N, Walch A, Koster RW (2009) Cadherin-2 controls directional chain migration of cerebellar granule neurons. PLoS Biol 7:e1000240PubMedPubMedCentralCrossRefGoogle Scholar
  210. Rivero S, Cardenas J, Bornens M, Rios RM (2009) Microtubule nucleation at the cis-side of the Golgi apparatus requires AKAP450 and GM130. EMBO J 28:1016–1028PubMedPubMedCentralCrossRefGoogle Scholar
  211. Rochlin MW, Wickline KM, Bridgman PC (1996) Microtubule stability decreases axon elongation but not axoplasm production. J Neurosci Off J Soc Neurosci 16:3236–3246Google Scholar
  212. Rodriguez OC, Schaefer AW, Mandato CA, Forscher P, Bement WM, Waterman-Storer CM (2003) Conserved microtubule-actin interactions in cell movement and morphogenesis. Nat Cell Biol 5:599–609PubMedCrossRefGoogle Scholar
  213. Rogers SL, Wiedemann U, Hacker U, Turck C, Vale RD (2004) Drosophila RhoGEF2 associates with microtubule plus ends in an EB1-dependent manner. Curr Biol CB 14:1827–1833PubMedCrossRefGoogle Scholar
  214. Roll-Mecak A, Vale RD (2008) Structural basis of microtubule severing by the hereditary spastic paraplegia protein spastin. Nature 451:363–367PubMedPubMedCentralCrossRefGoogle Scholar
  215. Rooney C, White G, Nazgiewicz A, Woodcock SA, Anderson KI, Ballestrem C, Malliri A (2010) The Rac activator STEF (Tiam2) regulates cell migration by microtubule-mediated focal adhesion disassembly. EMBO Rep 11:292–298PubMedPubMedCentralCrossRefGoogle Scholar
  216. Saillour Y, Broix L, Bruel-Jungerman E, Lebrun N, Muraca G, Rucci J, Poirier K, Belvindrah R, Francis F, Chelly J (2014) Beta tubulin isoforms are not interchangeable for rescuing impaired radial migration due to Tubb3 knockdown. Hum Mol Genet 23:1516–1526PubMedCrossRefGoogle Scholar
  217. Sakakibara A, Sato T, Ando R, Noguchi N, Masaoka M, Miyata T (2014) Dynamics of centrosome translocation and microtubule organization in neocortical neurons during distinct modes of polarization. Cereb Cortex 24:1301–1310PubMedCrossRefGoogle Scholar
  218. Sanz-Moreno V, Marshall CJ (2010) The plasticity of cytoskeletal dynamics underlying neoplastic cell migration. Curr Opin Cell Biol 22:690–696PubMedCrossRefGoogle Scholar
  219. Sapir T, Frotscher M, Levy T, Mandelkow EM, Reiner O (2012) Tau’s role in the developing brain: implications for intellectual disability. Hum Mol Genet 21:1681–1692PubMedCrossRefGoogle Scholar
  220. Schober JM, Cain JM, Komarova YA, Borisy GG (2009) Migration and actin protrusion in melanoma cells are regulated by EB1 protein. Cancer Lett 284:30–36PubMedCrossRefGoogle Scholar
  221. Schreiber SC, Giehl K, Kastilan C, Hasel C, Muhlenhoff M, Adler G, Wedlich D, Menke A (2008) Polysialylated NCAM represses E-cadherin-mediated cell-cell adhesion in pancreatic tumor cells. Gastroenterology 134:1555–1566PubMedCrossRefGoogle Scholar
  222. Schwartz MA (2001) Integrin signaling revisited. Trends Cell Biol 11:466–470PubMedCrossRefGoogle Scholar
  223. Shieh JC, Schaar BT, Srinivasan K, Brodsky FM, McConnell SK (2011) Endocytosis regulates cell soma translocation and the distribution of adhesion proteins in migrating neurons. PLoS One 6:e17802PubMedPubMedCentralCrossRefGoogle Scholar
  224. Shih W, Yamada S (2012) N-cadherin-mediated cell-cell adhesion promotes cell migration in a three-dimensional matrix. J Cell Sci 125:3661–3670PubMedPubMedCentralCrossRefGoogle Scholar
  225. Siegrist SE, Doe CQ (2007) Microtubule-induced cortical cell polarity. Genes Dev 21:483–496PubMedCrossRefGoogle Scholar
  226. Sirajuddin M, Rice LM, Vale RD (2014) Regulation of microtubule motors by tubulin isotypes and post-translational modifications. Nat Cell Biol 16:335–344PubMedPubMedCentralCrossRefGoogle Scholar
  227. Sit ST, Manser E (2011) Rho GTPases and their role in organizing the actin cytoskeleton. J Cell Sci 124:679–683PubMedCrossRefGoogle Scholar
  228. Small JV, Kaverina I (2003) Microtubules meet substrate adhesions to arrange cell polarity. Curr Opin Cell Biol 15:40–47PubMedCrossRefGoogle Scholar
  229. Spiczka KS, Yeaman C (2008) Ral-regulated interaction between Sec5 and paxillin targets Exocyst to focal complexes during cell migration. J Cell Sci 121:2880–2891PubMedPubMedCentralCrossRefGoogle Scholar
  230. Steffen A, Le Dez G, Poincloux R, Recchi C, Nassoy P, Rottner K, Galli T, Chavrier P (2008) MT1-MMP-dependent invasion is regulated by TI-VAMP/VAMP7. Curr Biol CB 18:926–931PubMedCrossRefGoogle Scholar
  231. Stehbens S, Wittmann T (2012) Targeting and transport: how microtubules control focal adhesion dynamics. J Cell Biol 198:481–489PubMedPubMedCentralCrossRefGoogle Scholar
  232. Stehbens SJ, Paterson AD, Crampton MS, Shewan AM, Ferguson C, Akhmanova A, Parton RG, Yap AS (2006) Dynamic microtubules regulate the local concentration of E-cadherin at cell-cell contacts. J Cell Sci 119:1801–1811PubMedCrossRefGoogle Scholar
  233. Steinecke A, Gampe C, Valkova C, Kaether C, Bolz J (2012) Disrupted-in-Schizophrenia 1 (DISC1) is necessary for the correct migration of cortical interneurons. J Neurosci Off J Soc Neurosci 32:738–745CrossRefGoogle Scholar
  234. Stiess M, Maghelli N, Kapitein LC, Gomis-Ruth S, Wilsch-Brauninger M, Hoogenraad CC, Tolic-Norrelykke IM, Bradke F (2010) Axon extension occurs independently of centrosomal microtubule nucleation. Science 327:704–707PubMedCrossRefGoogle Scholar
  235. Stramer B, Moreira S, Millard T, Evans I, Huang CY, Sabet O, Milner M, Dunn G, Martin P, Wood W (2010) Clasp-mediated microtubule bundling regulates persistent motility and contact repulsion in Drosophila macrophages in vivo. J Cell Biol 189:681–689PubMedPubMedCentralCrossRefGoogle Scholar
  236. Straube A (2011) How to measure microtubule dynamics? Methods Mol Biol 777:1–14PubMedCrossRefGoogle Scholar
  237. Straube A, Merdes A (2007) EB3 regulates microtubule dynamics at the cell cortex and is required for myoblast elongation and fusion. Curr Biol CB 17:1318–1325PubMedCrossRefGoogle Scholar
  238. Sudo H, Baas PW (2010) Acetylation of microtubules influences their sensitivity to severing by katanin in neurons and fibroblasts. J Neurosci Off J Soc Neurosci 30:7215–7226CrossRefGoogle Scholar
  239. Sudo H, Baas PW (2011) Strategies for diminishing katanin-based loss of microtubules in tauopathic neurodegenerative diseases. Hum Mol Genet 20:763–778PubMedPubMedCentralCrossRefGoogle Scholar
  240. Sudo H, Maru Y (2008) LAPSER1/LZTS2: a pluripotent tumor suppressor linked to the inhibition of katanin-mediated microtubule severing. Hum Mol Genet 17:2524–2540PubMedCrossRefGoogle Scholar
  241. Sun X, Li F, Dong B, Suo S, Liu M, Li D, Zhou J (2013) Regulation of tumor angiogenesis by the microtubule-binding protein CLIP-170. Protein Cell 4:266–276PubMedCrossRefGoogle Scholar
  242. Suzuki A, Ohno S (2006) The PAR-aPKC system: lessons in polarity. J Cell Sci 119:979–987PubMedCrossRefGoogle Scholar
  243. Takesono A, Heasman SJ, Wojciak-Stothard B, Garg R, Ridley AJ (2010) Microtubules regulate migratory polarity through Rho/ROCK signaling in T cells. PLoS One 5:e8774PubMedPubMedCentralCrossRefGoogle Scholar
  244. Takino T, Watanabe Y, Matsui M, Miyamori H, Kudo T, Seiki M, Sato H (2006) Membrane-type 1 matrix metalloproteinase modulates focal adhesion stability and cell migration. Exp Cell Res 312:1381–1389PubMedCrossRefGoogle Scholar
  245. Tanaka E, Ho T, Kirschner MW (1995) The role of microtubule dynamics in growth cone motility and axonal growth. J Cell Biol 128:139–155PubMedCrossRefGoogle Scholar
  246. Tassan JP, Le Goff X (2004) An overview of the KIN1/PAR-1/MARK kinase family. Biol Cell Under Auspices Eur Cell Biol Org 96:193–199Google Scholar
  247. Theisen U, Straube E, Straube A (2012) Directional persistence of migrating cells requires Kif1C-mediated stabilization of trailing adhesions. Dev Cell 23:1153–1166PubMedCrossRefGoogle Scholar
  248. Tobin JL, Di Franco M, Eichers E, May-Simera H, Garcia M, Yan J, Quinlan R, Justice MJ, Hennekam RC, Briscoe J, Tada M, Mayor R, Burns AJ, Lupski JR, Hammond P, Beales PL (2008) Inhibition of neural crest migration underlies craniofacial dysmorphology and Hirschsprung’s disease in Bardet-Biedl syndrome. Proc Natl Acad Sci U S A 105:6714–6719PubMedPubMedCentralCrossRefGoogle Scholar
  249. Toyo-Oka K, Sasaki S, Yano Y, Mori D, Kobayashi T, Toyoshima YY, Tokuoka SM, Ishii S, Shimizu T, Muramatsu M, Hiraiwa N, Yoshiki A, Wynshaw-Boris A, Hirotsune S (2005) Recruitment of katanin p60 by phosphorylated NDEL1, an LIS1 interacting protein, is essential for mitotic cell division and neuronal migration. Hum Mol Genet 14:3113–3128PubMedCrossRefGoogle Scholar
  250. Tsai FC, Seki A, Yang HW, Hayer A, Carrasco S, Malmersjo S, Meyer T (2014) A polarized Ca2+, diacylglycerol and STIM1 signalling system regulates directed cell migration. Nat Cell Biol 16:133–144PubMedPubMedCentralCrossRefGoogle Scholar
  251. Tsai JW, Bremner KH, Vallee RB (2007) Dual subcellular roles for LIS1 and dynein in radial neuronal migration in live brain tissue. Nat Neurosci 10:970–979PubMedCrossRefGoogle Scholar
  252. Tsai L-H, Gleeson JG (2005) Nucleokinesis in neuronal migration. Neuron 46:383–388PubMedCrossRefGoogle Scholar
  253. Tsvetkov AS, Samsonov A, Akhmanova A, Galjart N, Popov SV (2007) Microtubule-binding proteins CLASP1 and CLASP2 interact with actin filaments. Cell Motil Cytoskeleton 64:519–530PubMedCrossRefGoogle Scholar
  254. Umeshima H, Hirano T, Kengaku M (2007) Microtubule-based nuclear movement occurs independently of centrosome positioning in migrating neurons. Proc Natl Acad Sci U S A 104:16182–16187PubMedPubMedCentralCrossRefGoogle Scholar
  255. van der Vaart B, Akhmanova A, Straube A (2009) Regulation of microtubule dynamic instability. Biochem Soc Trans 37:1007–1013PubMedCrossRefGoogle Scholar
  256. van Es JH, Giles RH, Clevers HC (2001) The many faces of the tumor suppressor gene APC. Exp Cell Res 264:126–134PubMedCrossRefGoogle Scholar
  257. Van Haastert PJ, Devreotes PN (2004) Chemotaxis: signalling the way forward. Nat Rev Mol Cell Biol 5:626–634PubMedCrossRefGoogle Scholar
  258. van Haren J, Boudeau J, Schmidt S, Basu S, Liu Z, Lammers D, Demmers J, Benhari J, Grosveld F, Debant A, Galjart N (2014) Dynamic microtubules catalyze formation of navigator-TRIO complexes to regulate neurite extension. Curr Biol CB 24:1778–1785PubMedCrossRefGoogle Scholar
  259. Vasiliev JM, Gelfand IM, Domnina LV, Ivanova OY, Komm SG, Olshevskaja LV (1970) Effect of colcemid on the locomotory behaviour of fibroblasts. J Embryol Exp Morphol 24:625–640PubMedGoogle Scholar
  260. Vicente-Manzanares M, Ma X, Adelstein RS, Horwitz AR (2009) Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat Rev Mol Cell Biol 10:778–790PubMedPubMedCentralCrossRefGoogle Scholar
  261. Vicente-Manzanares M, Zareno J, Whitmore L, Choi CK, Horwitz AF (2007) Regulation of protrusion, adhesion dynamics, and polarity by myosins IIA and IIB in migrating cells. J Cell Biol 176:573–580PubMedPubMedCentralCrossRefGoogle Scholar
  262. Vinogradova T, Paul R, Grimaldi AD, Loncarek J, Miller PM, Yampolsky D, Magidson V, Khodjakov A, Mogilner A, Kaverina I (2012) Concerted effort of centrosomal and Golgi-derived microtubules is required for proper Golgi complex assembly but not for maintenance. Mol Biol Cell 23:820–833PubMedPubMedCentralCrossRefGoogle Scholar
  263. Vitriol EA, Zheng JQ (2012) Growth cone travel in space and time: the cellular ensemble of cytoskeleton, adhesion, and membrane. Neuron 73:1068–1081PubMedPubMedCentralCrossRefGoogle Scholar
  264. Vogl T, Ludwig S, Goebeler M, Strey A, Thorey IS, Reichelt R, Foell D, Gerke V, Manitz MP, Nacken W, Werner S, Sorg C, Roth J (2004) MRP8 and MRP14 control microtubule reorganization during transendothelial migration of phagocytes. Blood 104:4260–4268PubMedCrossRefGoogle Scholar
  265. Wakelam MJ (1985) The fusion of myoblasts. Biochem J 228:1–12PubMedPubMedCentralCrossRefGoogle Scholar
  266. Wang CQ, Qu X, Zhang XY, Zhou CJ, Liu GX, Dong ZQ, Wei FC, Sun SZ (2010) Overexpression of Kif2a promotes the progression and metastasis of squamous cell carcinoma of the oral tongue. Oral Oncol 46:65–69PubMedCrossRefGoogle Scholar
  267. Wang J, Ma S, Ma R, Qu X, Liu W, Lv C, Zhao S, Gong Y (2014) KIF2A silencing inhibits the proliferation and migration of breast cancer cells and correlates with unfavorable prognosis in breast cancer. BMC Cancer 14:461PubMedPubMedCentralCrossRefGoogle Scholar
  268. Wang Y, McNiven MA (2012) Invasive matrix degradation at focal adhesions occurs via protease recruitment by a FAK-p130Cas complex. J Cell Biol 196:375–385PubMedPubMedCentralCrossRefGoogle Scholar
  269. Watanabe T, Noritake J, Kakeno M, Matsui T, Harada T, Wang S, Itoh N, Sato K, Matsuzawa K, Iwamatsu A, Galjart N, Kaibuchi K (2009a) Phosphorylation of CLASP2 by GSK-3beta regulates its interaction with IQGAP1, EB1 and microtubules. J Cell Sci 122:2969–2979PubMedCrossRefGoogle Scholar
  270. Watanabe T, Sato K, Kaibuchi K (2009b) Cadherin-mediated intercellular adhesion and signaling cascades involving small GTPases. Cold Spring Harb Perspect Biol 1:a003020PubMedPubMedCentralCrossRefGoogle Scholar
  271. Watanabe T, Wang S, Noritake J, Sato K, Fukata M, Takefuji M, Nakagawa M, Izumi N, Akiyama T, Kaibuchi K (2004) Interaction with IQGAP1 links APC to Rac1, Cdc42, and actin filaments during cell polarization and migration. Dev Cell 7:871–883PubMedCrossRefGoogle Scholar
  272. Waterman-Storer CM, Salmon ED (1997) Actomyosin-based retrograde flow of microtubules in the lamella of migrating epithelial cells influences microtubule dynamic instability and turnover and is associated with microtubule breakage and treadmilling. J Cell Biol 139:417–434PubMedPubMedCentralCrossRefGoogle Scholar
  273. Waterman-Storer CM, Salmon ED (1998) Endoplasmic reticulum membrane tubules are distributed by microtubules in living cells using three distinct mechanisms. Curr Biol CB 8:798–806PubMedCrossRefGoogle Scholar
  274. Waterman-Storer CM, Salmon WC, Salmon ED (2000) Feedback interactions between cell-cell adherens junctions and cytoskeletal dynamics in newt lung epithelial cells. Mol Biol Cell 11:2471–2483PubMedPubMedCentralCrossRefGoogle Scholar
  275. Waterman-Storer CM, Worthylake RA, Liu BP, Burridge K, Salmon ED (1999) Microtubule growth activates Rac1 to promote lamellipodial protrusion in fibroblasts. Nat Cell Biol 1:45–50PubMedCrossRefGoogle Scholar
  276. Webb DJ, Donais K, Whitmore LA, Thomas SM, Turner CE, Parsons JT, Horwitz AF (2004) FAK-Src signalling through paxillin, ERK and MLCK regulates adhesion disassembly. Nat Cell Biol 6:154–161PubMedCrossRefGoogle Scholar
  277. Weber GF, Bjerke MA, DeSimone DW (2012) A mechanoresponsive cadherin-keratin complex directs polarized protrusive behavior and collective cell migration. Dev Cell 22:104–115PubMedPubMedCentralCrossRefGoogle Scholar
  278. Wen Y, Eng CH, Schmoranzer J, Cabrera-Poch N, Morris EJ, Chen M, Wallar BJ, Alberts AS, Gundersen GG (2004) EB1 and APC bind to mDia to stabilize microtubules downstream of Rho and promote cell migration. Nat Cell Biol 6:820–830PubMedCrossRefGoogle Scholar
  279. Werr J, Xie X, Hedqvist P, Ruoslahti E, Lindbom L (1998) beta1 integrins are critically involved in neutrophil locomotion in extravascular tissue In vivo. J Exp Med 187:2091–2096PubMedPubMedCentralCrossRefGoogle Scholar
  280. Wickstrom SA, Lange A, Hess MW, Polleux J, Spatz JP, Kruger M, Pfaller K, Lambacher A, Bloch W, Mann M, Huber LA, Fassler R (2010) Integrin-linked kinase controls microtubule dynamics required for plasma membrane targeting of caveolae. Dev Cell 19:574–588PubMedPubMedCentralCrossRefGoogle Scholar
  281. Wiesner C, Faix J, Himmel M, Bentzien F, Linder S (2010) KIF5B and KIF3A/KIF3B kinesins drive MT1-MMP surface exposure, CD44 shedding, and extracellular matrix degradation in primary macrophages. Blood 116:1559–1569PubMedCrossRefGoogle Scholar
  282. Willemsen MH, Vissers LE, Willemsen MA, van Bon BW, Kroes T, de Ligt J, de Vries BB, Schoots J, Lugtenberg D, Hamel BC, van Bokhoven H, Brunner HG, Veltman JA, Kleefstra T (2012) Mutations in DYNC1H1 cause severe intellectual disability with neuronal migration defects. J Med Genet 49:179–183PubMedCrossRefGoogle Scholar
  283. Williamson T, Gordon-Weeks PR, Schachner M, Taylor J (1996) Microtubule reorganization is obligatory for growth cone turning. Proc Natl Acad Sci U S A 93:15221–15226PubMedPubMedCentralCrossRefGoogle Scholar
  284. Wittmann T, Bokoch GM, Waterman-Storer CM (2004) Regulation of microtubule destabilizing activity of Op18/stathmin downstream of Rac1. J Biol Chem 279:6196–6203PubMedCrossRefGoogle Scholar
  285. Wolf K, Mazo I, Leung H, Engelke K, von Andrian UH, Deryugina EI, Strongin AY, Brocker EB, Friedl P (2003a) Compensation mechanism in tumor cell migration: mesenchymal-amoeboid transition after blocking of pericellular proteolysis. J Cell Biol 160:267–277PubMedPubMedCentralCrossRefGoogle Scholar
  286. Wolf K, Muller R, Borgmann S, Brocker EB, Friedl P (2003b) Amoeboid shape change and contact guidance: T-lymphocyte crawling through fibrillar collagen is independent of matrix remodeling by MMPs and other proteases. Blood 102:3262–3269PubMedCrossRefGoogle Scholar
  287. Wu X, Kodama A, Fuchs E (2008) ACF7 regulates cytoskeletal-focal adhesion dynamics and migration and has ATPase activity. Cell 135:137–148PubMedPubMedCentralCrossRefGoogle Scholar
  288. Wu Y, Song SW, Sun J, Bruner JM, Fuller GN, Zhang W (2010) IIp45 inhibits cell migration through inhibition of HDAC6. J Biol Chem 285:3554–3560PubMedPubMedCentralCrossRefGoogle Scholar
  289. Xie Z, Sanada K, Samuels BA, Shih H, Tsai LH (2003) Serine 732 phosphorylation of FAK by Cdk5 is important for microtubule organization, nuclear movement, and neuronal migration. Cell 114:469–482PubMedCrossRefGoogle Scholar
  290. Xu J, Wang F, Van Keymeulen A, Rentel M, Bourne HR (2005) Neutrophil microtubules suppress polarity and enhance directional migration. Proc Natl Acad Sci U S A 102:6884–6889PubMedPubMedCentralCrossRefGoogle Scholar
  291. Yadav S, Puri S, Linstedt AD (2009) A primary role for Golgi positioning in directed secretion, cell polarity, and wound healing. Mol Biol Cell 20:1728–1736PubMedPubMedCentralCrossRefGoogle Scholar
  292. Yam PT, Wilson CA, Ji L, Hebert B, Barnhart EL, Dye NA, Wiseman PW, Danuser G, Theriot JA (2007) Actin-myosin network reorganization breaks symmetry at the cell rear to spontaneously initiate polarized cell motility. J Cell Biol 178:1207–1221PubMedPubMedCentralCrossRefGoogle Scholar
  293. Yanagisawa M, Kaverina IN, Wang A, Fujita Y, Reynolds AB, Anastasiadis PZ (2004) A novel interaction between kinesin and p120 modulates p120 localization and function. J Biol Chem 279:9512–9521PubMedCrossRefGoogle Scholar
  294. Yau KW, van Beuningen SF, Cunha-Ferreira I, Cloin BM, van Battum EY, Will L, Schatzle P, Tas RP, van Krugten J, Katrukha EA, Jiang K, Wulf PS, Mikhaylova M, Harterink M, Pasterkamp RJ, Akhmanova A, Kapitein LC, Hoogenraad CC (2014) Microtubule minus-end binding protein CAMSAP2 controls axon specification and dendrite development. Neuron 82:1058–1073PubMedCrossRefGoogle Scholar
  295. Ye X, Lee YC, Choueiri M, Chu K, Huang CF, Tsai WW, Kobayashi R, Logothetis CJ, Yu-Lee LY, Lin SH (2012) Aberrant expression of katanin p60 in prostate cancer bone metastasis. Prostate 72:291–300PubMedPubMedCentralCrossRefGoogle Scholar
  296. Yilmaz M, Christofori G (2009) EMT, the cytoskeleton, and cancer cell invasion. Cancer Metastasis Rev 28:15–33PubMedCrossRefGoogle Scholar
  297. Yoo SK, Lam PY, Eichelberg MR, Zasadil L, Bement WM, Huttenlocher A (2012) The role of microtubules in neutrophil polarity and migration in live zebrafish. J Cell Sci 125:5702–5710PubMedPubMedCentralCrossRefGoogle Scholar
  298. Yu W, Qiang L, Solowska JM, Karabay A, Korulu S, Baas PW (2008) The microtubule-severing proteins spastin and katanin participate differently in the formation of axonal branches. Mol Biol Cell 19:1485–1498PubMedPubMedCentralCrossRefGoogle Scholar
  299. Yvon AM, Walker JW, Danowski B, Fagerstrom C, Khodjakov A, Wadsworth P (2002) Centrosome reorientation in wound-edge cells is cell type specific. Mol Biol Cell 13:1871–1880PubMedPubMedCentralCrossRefGoogle Scholar
  300. Zaidel-Bar R, Itzkovitz S, Ma’ayan A, Iyengar R, Geiger B (2007) Functional atlas of the integrin adhesome. Nat Cell Biol 9:858–867PubMedPubMedCentralCrossRefGoogle Scholar
  301. Zhang D, Grode KD, Stewman SF, Diaz-Valencia JD, Liebling E, Rath U, Riera T, Currie JD, Buster DW, Asenjo AB, Sosa HJ, Ross JL, Ma A, Rogers SL, Sharp DJ (2011) Drosophila katanin is a microtubule depolymerase that regulates cortical-microtubule plus-end interactions and cell migration. Nat Cell Biol 13:361–370PubMedPubMedCentralCrossRefGoogle Scholar
  302. Zhang Y, Chen K, Tu Y, Wu C (2004) Distinct roles of two structurally closely related focal adhesion proteins, alpha-parvins and beta-parvins, in regulation of cell morphology and survival. J Biol Chem 279:41695–41705PubMedCrossRefGoogle Scholar
  303. Zhou FQ, Zhou J, Dedhar S, Wu YH, Snider WD (2004) NGF-induced axon growth is mediated by localized inactivation of GSK-3beta and functions of the microtubule plus end binding protein APC. Neuron 42:897–912PubMedCrossRefGoogle Scholar

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© Springer-Verlag Wien 2016

Authors and Affiliations

  1. 1.Cellular and Molecular Neurobiology, Institute of ZoologyTU BraunschweigBraunschweigGermany
  2. 2.Cytoskeletal Dynamics, Centre for Mechanochemical Cell Biology, Warwick Medical SchoolUniversity of WarwickCoventryUK

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