No retraction, correction or concern found when last checked on Oct 18, 2026.

The Cytoskeleton—A Complex Interacting Meshwork

Tim Hohmann, Faramarz Dehghani

Cells · 2019 · Review · CC BY

Abstract

The cytoskeleton of animal cells is one of the most complicated and functionally versatile structures, involved in processes such as endocytosis, cell division, intra-cellular transport, motility, force transmission, reaction to external forces, adhesion and preservation, and adaptation of cell shape. These functions are mediated by three classical cytoskeletal filament types, as follows: Actin, microtubules, and intermediate filaments. The named filaments form a network that is highly structured and dynamic, responding to external and internal cues with a quick reorganization that is orchestrated on the time scale of minutes and has to be tightly regulated. Especially in brain tumors, the cytoskeleton plays an important role in spreading and migration of tumor cells. As the cytoskeletal organization and regulation is complex and many-faceted, this review aims to summarize the findings about cytoskeletal filament types, including substructures formed by them, such as lamellipodia, stress fibers, and interactions between intermediate filaments, microtubules and actin. Additionally, crucial regulatory aspects of the cytoskeletal filaments and the formed substructures are discussed and integrated into the concepts of cell motility. Even though little is known about the impact of cytoskeletal alterations on the progress of glioma, a final point discussed will be the impact of established cytoskeletal alterations in the cellular behavior and invasion of glioma.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 18, 2026. Paragraph numbers are ours, so an answer can point at one.

1. Introduction

1A single animal cell has the ability to adapt its shape in response to environmental confinements or chemical cues, to move through tissues (artificial and in vivo, including narrow spaces), and to divide. These processes are all, at least in part, orchestrated by the (dis-)assembly of cytoskeletal proteins. The cytoskeleton is made up of three major types of proteins, as follows: Tubulin, actin, and proteins forming intermediate filaments. These cytoskeletal proteins differ not only in their chemical structure, but also in the type of filaments and structures they form, ranging from fast assembling dendritic actin networks of the lamellipodium, capable of generating forces necessary for cell movement over single microtubule filaments as transport structures, to intermediate filaments capable of promoting or inhibiting cell movement and stabilizing the cell against large stress.

2To form a myriad of different cytoskeletal structures, as it is observed in animal cells, the cytoskeletal meshwork does not only need different components with different properties and functions, but also a tight and precise regulation of (dis-)assembly of its components, the respective local regulation of (dis-)assembly-factors, and interactions between the actin, microtubule, and intermediate filament networks.

3The ability of cells to migrate is of special interest in glioma spreading, as the success of glioma treatment is crucially coupled to the question of whether a recurrent tumor will arise or not, as resection is successful only if the tumor is completely removed. Consequently, recurrent tumor formation is considered to be the main reason of tumor morbidity. Hence, targeting the migratory machinery in gliomas can be a promising approach for the containment of metastasis. For successful targeting of glioma migration, a broad and detailed knowledge of the cytoskeletal architecture and its alterations is necessary.

4Here we provide an overview of different cytoskeletal filaments, including actin, microtubules, and intermediate filaments, their (dis-)assembly, interactions, and function in motility and shape changes of healthy cells. Afterwards, cytoskeletal alterations in glioma and their impact on their migratory behavior are discussed.

2. Actin Regulation and Structure

5In cells, actin occurs in two distinct states, as follows: The monomeric G-actin and filamentous F-actin. The modulation of the actin cytoskeleton is regulated by the balance of globular G- and polymeric F-actin and by actin associated proteins. The actin cytoskeleton forms a network consisting of polarized filaments that are mostly associated with force generation necessary for movement, focal adhesion, and shape changes. In the following we describe the building blocks of actin filaments, the assembly and disassembly of filaments, their kinetics, regulation, as well as filament bundles, network structures, and their mechanical properties. A summary of all mentioned actin binding or associated proteins and their function can be found in Table 1.

2.1. Actin Filaments

6Actin filaments are, in contrast to intermediate filaments and microtubules, semi-flexible filaments, forming dendritic or cross-linked structures. Semi-flexible means that the persistence length of a single filament is in the order of the filament length, where the persistence length is the length scale on which the correlation between two tangents along the filament drops to 1/e. As a semi-flexible polymer, actin filaments are actively bent by thermal fluctuations, thus generating additional resistance to forces stretching the filament. Actin itself is considered the most dynamic of the three cytoskeletal proteins capable of strong structural changes in the time scale of minutes, thus determining the shape of a cell. A single actin filament consists of actin monomers, called globular actin (G-actin). Under nearly physiological conditions G-actin polymerizes to asymmetric helical structures, filamentous actin (F-actin), with a typical length of 6–7 µm in in vitro studies. The nucleation kinetics is mostly limited by the generation of dimers and trimers. Having reached the trimer state, filament nucleation increases rapidly, but in dependence of the available G-actin pool (Figure 1). The resulting actin filaments have a right handed helical structure. G-actin is polarized, therefore F-actin is polarized as well, with the less dynamic side termed as the (−)-end and the more dynamic (+)-end having a ten times higher polymerization rate than the (−)-end. As actin is an ATPase the (+)- and (−)-ends can also be distinguished by their ATP/ADP status, especially if the growth at the (−)-end is inhibited further. Thus the (+)-end contains higher amounts of ATP bound actin while the (−)-end contains more ADP bound actin.

2.2. Profilin

7As F-actin is capable of forming spontaneously above a certain critical concentration of G-actin (≈0.1 µM) a precise cofactor-driven control of actin polymerization is necessary. One such control element is profilin. Profilin is an actin binding protein that regulates actin homeostasis by inhibiting the spontaneous formation of actin di- and trimers, but it also catalyzes the transition from ADP- to ATP-actin. Profilin bound G-actin can be used for the construction of actin filaments if nucleation factors like the Arp2/3 (actin-related-proteins 2/3) complex or formins are present. Interestingly, if formins and profilin are present, free actin elongation can be increased by a factor of up to 9. Additionally, profilin was shown to inhibit polymerization at the (−)-end of actin.

2.3. Dendritic Actin Networks

8Besides the quasi-linear actin filaments there are dendritic actin networks, formed by the Arp2/3 complex. These networks are usually formed at the cell front on a short time scale and, thus, its regulation is of high importance. The generation of a dendritic actin network starts from a so called primer, an existing actin filament at which Arp2/3 binds to its side. Arp2/3 is, amongst others, activated by members of the WASP family. For the generation of a dense dendritic network not only nucleation factors, but also capping proteins are needed to restrict the elongation of the actin (+)-ends. If capping proteins are present the Arp2/3 complex can generate multiple networks originating from different actin filaments that are able to merge and generate forces near the cell membrane. The number of nodes is important for the mechanical properties of the generated network and consequently the elastic modulus scales with the mesh size M by 1/M4. In general the dendritic network behaves visco-elastic, meaning it is mainly elastic on small time scales (<1 min) and viscous on longer time scales (>10 min).

2.4. Non-Muscle Myosin

9Another important molecule class is the myosin family. Here we will focus exclusively on the non-muscle myosin. Myosin is responsible for the contractility of anti-parallel actin structures using ATP hydrolysis as the energy supply. These contractile structures are mainly responsible for the retraction of the cell rear for productive movement, but also for transmitting forces to the surrounding extra-cellular matrix.

10Interestingly, myosin II motor activity alone is insufficient to produce contractility. Single myosin II hexamers are unipolar and thus ineffective in generating contractile forces, but when assembled into bipolar mini-filaments they are highly processive and capable of generating forces by pulling on anti-parallel actin filaments. Myosin II can be activated via phosphorylation of the regulatory light chain (RLC) or activation of myosin light chain kinases (MLCK). RLC is activated via Rho-associated protein kinase (ROCK) or citron kinases (both activated by RhoA) and MLCK by Ca2+. After RLC phosphorylation myosin is capable of generating contractile forces. Another type of regulation works via the phosphorylation of the myosin heavy chain, utilizing myosin heavy chain kinases (MHCK), casein kinase II (CKII), or protein kinase C (PKC), inhibiting mini-filament assembly or dissociating existing mini-filaments. The switch between those two activation states influences the contractility of the respective acto-myosin network. Consequently, regulation of myosin strongly impacts organization and properties of contractile actin structures, as discussed below.

2.5. Cross-Linked Actin Networks and Actin Bundles

11Despite the already mentioned dendritic actin structures, there are actin bundles and networks linked together by cross-linkers. Cross-linkers are molecules that connect single actin filaments either transiently or non-transiently and are either passive (e.g., scruin, fascin, α-actinin, filamin, or fimbrin) or active (myosin). Cross-linked actin bundles and networks largely control shape, mechanical integrity, and contractility of a cell. Generally, cross-linkers do not influence actin assembly (except for Arp2/3). Cross-linkers bind actin filaments based on their own size and the position of their binding-sites in different distances, ranging from 10 nm for fimbrin to 160 nm for filamin, and thus determine the density of the resulting actin structure. Additionally, the speed of actin polymerization influences the presence of cross-linkers in the resulting network, supposedly by crowding effects, thus excluding larger cross-linkers like α-actinin in quickly polymerizing filaments. If the formed actin structures are subject to a force that acts on a longer time scale than the binding time of the cross-linkers itself, a reorganization of cross-linkers and a subsequent shape change of the bundle occurs. This time scale depends on the type of cross-linker and its binding and unbinding time, which can be in the order of seconds. Consequently, cross-linked actin is elastic on short and viscous on long time scales and the presence of cross-linkers generally increases the elastic part of the visco-elastic answer to external stress.

12If actin filaments are bundled by cross-linkers the filaments inside the bundle can either be oriented in parallel or anti-parallel, meaning that (+)-ends of neighboring filaments are pointing in the same or the opposite direction. Parallel actin bundles are found amongst others in filopodia, while anti-parallel bundles are mostly found in stress fibers.

13Two mechanisms were proposed to explain the generation of parallel actin bundles. The first mechanism involves the Arp2/3 complex-dependent elongation of filaments in the absence of capping proteins, so that free growing (+)-ends transition into bundles via electro-static interactions between filaments. Here, the geometric constraints and the angle at which filaments make contact to each other determine whether parallel or anti-parallel bundles are formed. Notably, geometric constraints refer to all spatial limitations, such as the available free space, steric effects, etc., that potentially affect the final organization. Parallel bundles are then stabilized via cross-linkers, such as fascin. The second mechanism is formin dependent. Thereby the FH1 (formin homology) domain of formin functions as a ring structure capturing profilin bound actin molecules, while the FH2 domain interacts with the (+)-end of the filament. Some members of the formin family also move from the end of the filament into the middle, additionally functioning as cross-linkers to stabilize the generated structure. As formins do not necessarily bundle filaments, further proteins, such as cross-linkers or Ena/VASP, are also involved in formin dependent bundle formation. Ena/VASP is a protein family associated with anti-capping function and elongation, but has no nucleation activity on its own.

14In contrast to parallel bundles, anti-parallel bundles are mostly connected with classical cross-linkers and the motor-protein myosin, which has the ability to actively move antiparallel fibers relative to each other. As with parallel bundles, anti-parallel bundles are stabilized by cross-linkers favoring this configuration, like α-actinin or fimbrin. Through the activation of myosin anti-parallel bundles are pre-stressed, leading to either a contraction or dissociation of the bundle. Without further cross-linkers, anti-parallel bundles containing myosin first contract strongly and later disassemble.

15The mechanical properties of (anti-)parallel bundles depends on the type and density of cross-linkers and, thus, on the compactness of the bundle and whether the bundle allows the sliding of single filaments. For non-cross-linked bundles the persistence length scales with the number of filaments while for cross-linked bundles that allow no filament sliding it scales with the number of filaments squared.

2.6. ADF/Cofilin Induced Actin Disassembly

16As most of the actin structures are stable in time, cells need a mechanism to induce actin disassembly to adapt to environmental cues. One such mechanism is governed by the actin binding ADF/cofilin protein family, capable of disassembling and fragmenting actin filaments, but incapable of altering the polymerization rate. The efficiency of ADF/cofilin depends on its binding state to actin filaments. Filaments that are fully decorated with ADF/cofilin are stabilized while partially decorated filaments fragment faster. The induction of fragmentation is likely caused by a reduced persistence length of the filament, that can drop to ≈20% of its initial value through ADF/cofilin binding, locally generating increased mechanical stress. ADF/cofilin preferentially binds to the (−)-end of actin filaments (ADP bound actin), but binding to ATP bound actin favors its transition from ATP to ADP bound actin and, thus, accelerates filament dissociation. The preference for older ADP bound actin implies that ADF/cofilin disassembly mostly affects inactive compartments of the actin network. As ADF/cofilin cannot bind to the free (+)-end it can only fully saturate actin filaments if the (+)-end is bound by capping proteins and is not capable of further elongation. With the exception of fascin, cross-linkers and tension reduce the efficiency of ADF/cofilin fragmentation. Furthermore, ADF/cofilin is present in high concentrations at the leading edge in dendritic networks, fragmenting links generated by Arp2/3 and the actin filaments itself, generating more free (+)-ends.

2.7. Actin Structures Inside the Cell

17Looking at a motile cell the net movement is the result of multiple, mostly actin-dependent, processes, as follows: Formation of protrusions in direction of motion, subsequent adhesion to the substrate and loss of adhesion on the rear of the cell, followed by rear-contraction (reviewed in). These processes are governed by sub-cellular structures, like filopodia and the lamellipodium governing cell motion, while stress fibers and the cortex secure mechanical stability and contractility. Further types of protrusions are so called blebs, which are capable of regulating cell movement independently of filopodia and the lamellipodium. The interactions of actin discussed here and its structural and functional integration with microtubules and intermediate filaments are summarized in Figure 2.

2.7.1. The Lamellipodium

18The lamellipodium (Figure 3) is a flat structure mainly associated with cell movement, formed by the polymerization of actin at the cell front, while it is depolymerized at the back of the lamellipodium by ADF/cofilin refilling the G-actin pool. The continuing (de-)polymerization of the whole network creates a treadmilling effect and a retrograde actin flow in the cell, which is enhanced in some cell types by myosin induced depolymerization at the back of the lamellipodium. Any flow originating from the contraction of the rear via stress fibers generates a flow of opposite direction. The forces generated by actin polymerization in the lamellipodium are up to a few hundred pN/µm. The most important factor for the generation of the lamellipodium is the intrinsically inactive Arp2/3 complex that becomes activated by the Scar/WAVE complex in an activation process by the small Rho GTPase Rac1. Arp2/3 nucleates a new actin filament at the site of existing filaments. For a three dimensional environment, N-WASP (and not WAVE) was shown to activate Arp2/3 and Rac1 was not found to be strongly localized at the cell front. Actin growth is further promoted by the presence of members of the Ena/VASP family accumulating at the lamellipodial tip, promoting further actin elongation and preventing capping. Despite the active Arp2/3 complex, a capping protein is needed as well to limit the elongation of single filaments so they remain productive and do not form bundles with other uncapped filaments or buckle under the load. For the generation of a stable dendritic network, it is cross-linked by proteins such as cortactin. As the described regulation by Rac1 would result in a constant growth of the lamellipodium, it has to be restricted by a negative feedback loop. One possible mechanism is via the protein arpin, which inhibits Arp2/3 activity in the lamellipodium. It has been postulated that arpin is recruited by Rac1. Thus, it seems possible that Rac1 activation initiates lamellipodium growth via quick Arp2/3 recruitment and successive actin polymerization and later inhibits its growth via recruitment of arpin. A high turnover rate of arpin or significantly higher concentration might be necessary to inactivate Arp2/3. A proof for this kind of hypothesis is yet lacking. Despite the actin dynamics, the lamellipodium is also influenced by the cell membrane and its surface tension. A higher membrane surface tension led to a more oriented actin filament polymerization while a lower tension resulted in more protrusions, probably related to the finite forces generated by the lamellipodium. Regarding the mechanical properties of the lamellipodium, it has to be noted that myosin was observed to be present at the rear of the lamellipodium, explaining why the lamellipodium is elastic on short and viscous on long time scales.

19Due to Arp2/3 the actin in the lamellipodium is connected to a dendritic structure. Interestingly, an analysis of cell speed relative to the actin orientation in the lamellipodium could demonstrate that faster cells have filaments that are not exactly oriented in the direction of movement and the parallel orientation of filaments is associated with slower movement.

2.7.2. Filopodia

20Further structures associated with cell motility are filopodia (Figure 3). Filopodia are associated with a sensory function in neurons, but do not seem necessary for migration, as the fast moving corneal keratocytes do not possess filopodia in two dimensions and forces generated by filopodia are significantly smaller than those generated by the lamellipodium. In other systems there might be a role for filopodia in migration, e.g., in three dimensional systems. Filopodia form a structure consisting of parallel actin bundles, with their (+)-ends pointing in direction of the cell membrane. This orientation is established via formins (e.g., FMNL2) and Ena/VASP, both being capable of maintaining a prolonged actin polymerization. Some of these formins, like mDia2, can be activated by the small GTPase Cdc42. Cdc42 is also capable of activating N-WASP and thus Arp2/3, leading to filopodia formation. A common model for filopodia initiation suggests that actin polymerization occurs in the presence of activated Arp2/3 and without capping proteins forming actin bundles. Nevertheless, Arp2/3 does not seem to be necessary for filopodia initiation in adherent cells. A further model of filopodia initiation states that filopodia are initiated by clusters of activated formins near the plasma membrane, nucleating and elongating actin filaments. For both models, subsequent further elongation via formins (e.g., mDia2) and Ena/VASP and stabilization and bundling with cross-linkers, like fascin, generates “mature” filopodia. Besides their role in cell movement, filopodia initiate cell-cell contacts, transmit cell-cell-signals, and respond to the mechanical properties of their surroundings. Interestingly, when filopodia are retracted to the cell the myosins II,V, and VI are not involved in this process. This leads to the idea that only actin (de-)polymerization and changes in the cortex are responsible for filopodia dynamics.

2.7.3. Stress Fibers

21Another type of actin related structures are stress fibers (Figure 3) that are neither present in filopodia nor in the lamellipodium. Stress fibers are formed from bundles of anti-parallel actin filaments containing myosin II or parallel filaments. Stress fibers are assembled bundles of 10–30 actin filaments, cross-linked by α-actinin in a bipolar fashion, and linked to focal adhesions. Focal adhesions are binding sites that connect the cell to the substrate. Contractile stress fibers are one of the main contributors to cell contractility in many animal cells. As the contractility of these stress fibers is regulated by myosin, regulation of stress fiber contractility is in many ways similar to the regulation of myosin activity discussed before. In non-motile cells, stress fibers are usually thick and comparably stable, while motile cells typically contain fewer less pronounced fibers with a higher dynamic. Actin and myosin are the two principal constituents of contractile stress fibers, while non-contractile ones do not contain myosin. Despite these components, stress fibers contain actin binding proteins and focal adhesion-associated proteins binding and unbinding in quick succession. The molecules found in stress fibers include cross-linkers, such as α-actinin, which does not only function to stabilize the bundle but is also associated with kinases and signaling proteins and, thus, functioning as a signaling mediator. Stress fibers can contain further cross-linkers, like fascin, filamin, and paladin, but their precise role despite bundling remains elusive. One hypothesis states that these proteins function as basis for regulation of cytoskeletal dynamics as, for example, paladin interacts with profilin and VASP. Further molecules of e.g., the calponin, tropomyosin, caldesmon family, and others, are found in stress fibers and are all suggested to be part of the cytoskeletal and/or stress fiber regulation. Generally speaking, stress fiber formation has been directly associated with an activation of the formin mDia1 and the small Rho GTPase RhoA, activating ROCK. The formin favors prolonged actin polymerization of parallel filaments important for dorsal stress fibers. In contrast ROCK activates the LIM kinase (LIMK), which inhibits ADF/cofilin induced filament severing, and additionally, ROCK activates myosin, favoring stress fiber formation. Nevertheless, both the ROCK and formin mechanisms are necessary for the formation of contractile stress fibers. Two other Rho GTPases, Cdc42 and Rac1, act in more indirect ways via the induction of lamellipodial growth via Arp2/3 (Rac1) and filopodia formation via the formin mDia2 (Cdc42). Collapse of both filament types can function as seeds for either transversal or ventral stress fibers.

22Since stress fibers vary in their morphology, molecular signature, and association with focal adhesions, the four following types of stress fibers can be distinguished: The perinuclear actin cap, transverse arcs/stress fibers, and dorsal and ventral stress fibers.

23The three classes of contractile stress fibers are the ventral and transverse stress fibers and the perinuclear actin cap, all characterized by the presence of myosin II along the fibers. Even so, the myosin II spacing can change over time, indicating that contractile stress fibers are dynamic structures with non-uniform mechanical properties. Measurements indicate that stress fibers have a stiffness of roughly 12 kPa, constant for strains up to 0.12. Perturbation of myosin in stress fibers reduces the elastic modulus to 8 kPa, indicating the importance of myosin II in contractile stress fibers. Not surprisingly, the tensile elastic modulus increases from approximately 1.5 MPa to 104 MPa for strains approximating 2. Ventral stress fibers are oriented parallel to the direction of cell motion and connect adhesion sites of the cell, while transverse fibers are oriented perpendicular to the ventral fibers and are not directly connected to focal adhesions. Even so, transversal stress fibers can contribute to the overall contractility through their connection to dorsal stress fibers. The formation of transversal stress fibers is dependent on Arp2/3 and myosin and possess a periodic α-actinin-myosin pattern. Transversal stress fibers form when the dendritic network collapses and is restructured by myosin. Notably, simulations on the capability of myosin to generate contractile structures suggest that the presence of myosin and actin is sufficient to generate anti-parallel/contractile bundles, as these were found to be energetically favorable. A further origin of both transversal and ventral fibers is the collapse of filopodia, which functions as a seed for stress fibers. Additionally, ventral stress fibers can be formed from existing dorsal stress fibers and the attached transverse stress fibers, as well as by the fusion of two dorsal stress fibers. Ventral stress fibers are also contractile actin-myosin bundles attached to focal adhesions at both ends, thus being directly part of the contractile machinery. Due to their location at the rear of the cell and an orientation that is roughly in the direction of motion, they are part of the rear contraction, and thus associated with cell motility. The third type of contractile stress fibers is the so-called perinuclear cap, consisting of stress fibers positioned above the nucleus, regulating the shape of the nucleus. Additionally, they are proposed to serve as a mechanical connection between the nucleus and the rest of the cell. All of these contractile stress fiber types have in common that they are highly dependent on presence and activity of myosin and, thus, on tension. Consequently, myosin inhibition leads to the disassembly of these stress fibers.

24In contrast to the other stress fiber types, dorsal stress fibers do not contain myosin II and are anchored at focal adhesions at their distal ends. The lack of myosin directly leads to the lack of contractility of dorsal stress fibers. It is proposed that these fibers consist of fast growing (+)-ends that face the cell periphery and more distant parts consisting of mixed polarity actin filaments. Furthermore, paladin and Rac1 are seemingly essential for the formation of dorsal stress fibers. Paladin promotes fiber assembly via VASP recruitment. Functionally, these stress fibers seem to be an anchor point for the assembly of the other stress fiber types and a link to focal adhesions. It is supposed that dorsal stress fibers are generated via actin polymerization at emerging focal adhesions and stabilized during retraction phases of the lamellipodium via coupling to emerging transverse stress fibers.

2.7.4. Actin Cortex and Blebs

25The last cytoplasmic structure described here is the actin cortex (Figure 3), which forms a contractile actin structure at the boarder to the plasma membrane. The cortex is a few hundred nanometer thick layer, consisting of a mixture of filament bundles and cross-linked filaments, with a mesh-size of approximately ≈50–150 nm, a thickness of 50–100 nm, and a distance to the cell membrane of less than 20 nm. The cortex meshwork appears to be mainly isotropic, oriented in parallel to the plasma membrane, but some filaments are also oriented perpendicular to the membrane. Despite actin filaments, the cortex contains a number of cross-linkers (e.g., fascin, actinin, filamin, etc.), myosin, proteins that control actin turnover (like profilin, cofilin), capping proteins, proteins of the ERM (ezrin, radixin, moesin) family, nucleating factors (like Arp2/3, the formin mDia1), and signaling molecules such as RhoGTPases, RhoGEFs (guanine exchange factors), and RhoGAPs (GTPase activating proteins). The two mentioned nucleating factors Arp2/3 and mDia1 were also found to be responsible for the majority of cortical F-actin generation, while the ERM proteins link the cortex to the membrane and can therewith transmit forces acting on the membrane and determine the cell shape. Depletion of cofilin-1 or capping proteins in HeLa cells increased cortex thickness but reduced tension, implying a role for actin regulating proteins in cortical tension. Mechanical properties of the cortex determine how the cell deforms in response to external forces. On time scales smaller than the remodulation time of the cortex it behaves elastic, with a cell type-dependent elastic modulus in the order of a few hundred to thousands of pascals. On long time scales (>1min) the cortex behaves viscous because of the adaption to external forces via actin modulation, dissociation, and (un-)binding of cross-linkers. If myosin is activated the cortex turnover times can be even lower, perhaps via direct disassembly or enhanced actin breakage. Generally speaking, the behavior of the cortex is similar to that of glassy materials and consistent with relaxations of three dimensional in vitro actin networks.

26One of the main global and local properties of the cortex is its tension, which regulates the cell shape of single cells and tissues. Several studies demonstrated that the cortex tension depends on the myosin activity and actin polymerization, with higher myosin activity and lower actin polymerization leading to an increased cortex tension. A lower cortex tension is further associated with an increased protrusive activity of the cell, thus indirectly regulating cell motility. Interestingly, local drops in cortex tension or cortex-membrane adhesion and local ruptures of the cortex can be the origin of so called blebs (Figure 3), a special, initially actin free, membrane protrusion. Blebs can be initiated by any type of cortex weakening or loss of cortex-membrane adhesion if a given internal hydrostatic pressure threshold is reached. Localized myosin contractions, promoting either cortex tearing or increasing local intracellular pressure, are some of the main sources of blebbing, but others are also discussed. Thus, the activation of myosin via the already described activation by ROCK or MLCK are sufficient to induce bleb formation. The progress of a bleb can be divided in three steps, as follows: Initiation, growth, and retraction. Initially, the growing bleb does not contain actin, but over time, when the bleb expands further, the actin cortex reassembles at the plasma membrane, stalling the bleb growth up to the point of a full restoration where the generated contractile forces retract the bleb. It has to be noted that bleb retraction does not always occur and in some motile cells blebs are stabilized and used as an alternative or additional mode of migration. The expansion of blebs by actomyosin contraction induced pressures lasts 5–30 s, accompanied by a flow of cytosol into the bleb and a concomitant increase in surface area. The surface area is increased by a flow of lipids through the tearing of the membrane from the actin cortex. The maximal bleb size is determined by the initial growth rate and the cortex re-polymerization time, both being dependent on cortex tension. The concept of tension inhibiting bleb expansion is further supported by the idea that the needed membrane unfolding is effectively resisting bleb expansion and, thus, slowing down the growth. After full maturation the cortex is reconstructed and if the bleb is not stabilized via adhesions it is retracted by the re-established cortex via a myosin induced contraction.

2.7.5. Nuclear Actin

27For completeness it has to be mentioned that actin is not only present in the cytoplasm of eukaryotic cells, but also in the nucleus. As in the cytoplasm, nuclear actin exists in a monomeric and polymeric form. Nuclear G-actin was shown to associate with all three RNA polymerases, participating in transcription initiation and elongation. The exact function of G-actin in the transcription complex remains unclear, but G-actin levels need to be precisely tuned for normal translation and cofilin is required for its elongation. Contrary, stable actin filaments inhibit transcription. Furthermore, actin is implied to affect the nuclear structure. During nuclear expansion at the mitotic exit, chromatin reorganizes depending on the transient formation of polymeric actin, in a seemingly cofilin dependent manner. It is probable that nuclear actin regulates the structure of the nuclear envelope and the nucleus via interactions with the nuclear intermediate filaments lamin.

28As mentioned, cell nuclei contain polymeric actin that can be generated inside the nucleus via actin nucleation factors. These actin assembly factors include, amongst others, mDia1, Spire1/2, Fmn2, and Arp2/3. The presence of assembly factors underlines a functional role of polymeric actin and actin binding proteins. Initiation of DNA replication was demonstrated to require formin dependent nuclear actin polymerization. Additionally, the loss of Spire1/2 and Fmn2 resulted in a less efficient clearance of DNA double strand breaks. This is in agreement with two studies demonstrating an association of actin and Arp2/3 with sites of DNA damage and decreased damage repair after reduced Arp2/3 dependent actin nucleation. For more information on the role of actin in the nucleus the interested reader is referred to other reviews.

3. Microtubules

29Microtubules consist of α- and β-tubulin heterodimers forming hollow filaments, usually consisting of 13 protofilaments. The microtubules are seeded by the microtubule organization center (MTOC), generally centrosomal, but in some types of differentiated cell non-radial microtubules are assembled by non-centrosomal MTOCs, for example at the Golgi apparatus. MTOC, amongst others, contains γ-tubulin as a microtubule nucleator, as well as anchoring and adaptor proteins for attachment of microtubules. Heterodimers interact at the MTOC with the γ-TURC, thus microtubules are nucleated and anchored with their (−)-ends to the MTOC. Microtubules show a behavior called dynamic instability, characterized by a sudden switch from growing to growth arrest and/or quick depolymerization (termed catastrophe), followed by a new growth cycle (Figure 4). A possible explanation for this behavior can be derived from the formation process of single protofilaments. During polymerization, GTP-bound tubulin heterodimers are bound to the (+)-end and normally hydrolyzed shortly after, but sometimes older parts of microtubules still contain GTP-bound tubulin heterodimers. This leads to the following model: If, due to stochastic fluctuations or other perturbations, the (+)-end does not contain the more stable GTP-bound tubulin, it depolymerizes and is stabilized at older GTP-bound tubulin sites. Stochastic fluctuations in the rate of microtubule growth and the stochastic nature of GTP hydrolysis lead to a dynamic GTP cap size, with the consequence that faster polymerizing microtubules have a larger GTP cap, resulting in less frequent catastrophe events. The dynamic behavior of microtubules can be regulated by both intrinsic and extrinsic factors (see Table 2 for summary). Microtubule interacting proteins are either microtubule (+)-end-binding proteins (+TIP) or structural microtubule-associated proteins (MAP) interacting with microtubules along their length. These protein classes can have stabilizing or destabilizing effects, changing polymerization dynamics or severing microtubules. Important proteins belonging to the family of the +TIPs are CLASPs (cytoplasmic linker associated protein) and APC (adenomatous polyposis coli) that suppress microtubule catastrophe events and promote rescue after catastrophe. Part of the stabilizing effects of CLASPs arises from their capability to modulate interactions between microtubules and the cell cortex. Further important families of +TIPs are the spectraplakins, binding both microtubules and actin and EBs (end binding proteins). EBs are supposed to be a master regulator of +TIP recruitment (e.g., CLASP, APC, MACF1 (microtubule-actin crosslinking factor)) and complex assembly, generally promoting persistent microtubule growth. EBs are generally associated with an increased polymerization rate and reduced catastrophe number. Despite these molecules that mainly (de-)stabilize microtubules, there is a bunch of proteins that sever microtubules, like spastin or katanin, or influence depolymerization and polymerization, e.g., stathmin (favors depolymerization) or XMPA215 (increases polymerization rate). Additionally, there are structural MAPs, like tau protein, MAP2, or DCX (doublecortin), that interact with the filament at its whole length and stabilize it by reducing shrinkage speed, promoting filament growth, and reducing catastrophe frequency. The effect of structural MAPs can also inhibit the effect of other microtubule associated proteins, as, for example, tau protein can inhibit the katanin induced severing. A further important class of MAPs are the motor proteins kinesin and dynein, both serving as cargo transporters, exploiting the microtubule meshwork. In general, kinesin motor proteins transport cargo to the (+)-end, while dynein moves to the (−)-end of microtubules, transporting diverse cargo types, including membrane components, signaling molecules, such as the small GTPases Rac and Cdc42, but also intermediate filaments and their precursors, β-actin coding mRNA, and sub-units of the Arp2/3 complex. Motor proteins possess not only a transport function but can stabilize or destabilize microtubules. For example, members of the kinesin-8, kinesin-13 family, or KinI kinesins can induce depolymerization, likely via the destabilization of the GTP cap or the induction of kinks. Despite their transport function and regulatory role in microtubule dynamics, some kinesins organize the microtubule network via the bridging of microtubules, thus favoring the generation of parallel arrays. For example, in neurons kinesin-5 and kinesin-12 are necessary for axonal outgrowth because of their cross-linking ability and the concomitant focus on the extension of microtubule arrays. Additionally, kinesin-1 may be involved in this process by sliding filaments alongside each other. A further important aspect regulating microtubule dynamics are the post-translational modifications modifying microtubule properties and affinities of MAPs. Important modifications are, amongst others, tyrosination, glutamylation, and acetylation. Acetylation protects microtubules against repeated mechanical stress via an increased flexibility, but does not protect against depolymerization. Additionally, some severing proteins, such as katanin, preferentially interact with acetylated tubulin. Tyrosination affects the recruitment of microtubule interacting proteins, such as CLIP-170 or kinesin-1, that prefer detyrosinated microtubules, probably facilitating directional transport. In contrast, spastin favors cleavage of detyrosinated microtubules. Similar to tyrosination, glutamylation can also affect the interaction with microtubule associated proteins. Map2, tau, and kinesin-1 were reported to preferentially interact with those microtubules with up to three glutamates on their tail. Similarly, both microtubule severing proteins, katanin and spastin, show an increased affinity for glutamylated microtubules. Taken together, this data suggests that post translational modifications are important and finely tunable regulators of microtubule dynamics and, consequently, of cell behavior.

30For regulation of MAPs, and microtubules in general, the family of Rho GTPases is of major significance. An important example is the stabilizing effect of RhoA, but not Rac1 and Cdc42, on microtubules via the RhoA effector mDia. mDia is capable of interacting with EB1 and APC, leading to a stabilization of microtubules via e.g., Kif4. Furthermore, active mDia induces the alignment of actin and microtubules. Interestingly, stathmin action seems to negatively regulate the RhoA/ROCK activity, complementing the observation that Cdc42 and Rac1 phosphorylate stathmin via an activation of PAK (p21-activated kinases). In fact, Rac1 activation was demonstrated to decrease catastrophe events and increase microtubule growth time in cells via PAK. Furthermore, both Rac1 and Cdc42 activate IQGAP1, interacting with Clip-170, likely providing a stabilization site for microtubule (+)-ends near the cortex. The interactions of microtubules discussed here and their interactions with actin and intermediate filaments are summarized in Figure 2 and Figure 3.

31Microtubules typically play a role as tracks for transport, as already mentioned, in spindle positioning during mitosis, migration (discussed below), and in cell shape control. At the current point, the concept that microtubules control the balance between RhoA and Rac1 and thus influence cell shape and migration is favored over a direct mechanical participation for most cell types. Even so, microtubules are relatively stiff polymers, when compared to actin, capable of generating forces of up to 3–4 pN during polymerization. As a result, microtubules can deform membranes and resist compressional forces in such a way that they act as load bearing fibers in living cells via transversal re-enforcement by other cytoskeletal components. Notably, if multiple microtubule filaments grow as a bundle the generated forces increase linearly with the number of microtubules per bundle. Complementary to these observations, the load bearing capacities of microtubules is limited because compressional loads can induce catastrophe events, in line with the fact that most catastrophes are indeed induced at the cell edge and the observed short-wavelength buckles near the boundary of adherent cells. A further mechanism of force generation by microtubules is during the shrinkage phase. To actually transmit a force during shrinkage, the (+)-end needs to stay attached to its cargo. When the GTP cap is lost the microtubule protofilaments lose their lateral connection with neighboring protofilaments, bending backwards and forming ring-like shapes (Figure 4). If cargo stays attached during this process, a single microtubule can exert forces of up to 30–65 pN, a magnitude larger than the pushing force.

32Microtubules also play a key role during the separation of chromosomes during cell division. Depolymerization of microtubules is believed to generate the needed forces to separate the sister chromatids. This aspect will not be discussed in more detail here, but the interested reader may be referred to the following reviews:.

33Furthermore, microtubules and actin are linked regarding their functional dynamics and structural organization. On the one hand there is an indirect co-regulation, as microtubules are able to locally regulate and are regulated by RhoGTPases and focal adhesions (see also chapter 5.2), but on the other hand, there are molecules interacting with microtubules and actin. One such molecule is APC, which stabilizes microtubules and nucleates actin filaments, with actin nucleation additionally favored by the formin mDia1. A further formin mDia2 that is capable of nucleating actin can also stabilize microtubules, independent of its nucleation function. There are additional cross-linkers connecting actin and microtubule filaments, such as MACF1 and Arg. Consequently, the actin and microtubule cytoskeleton cannot fully be regarded as decoupled systems, as they are not regulated independently and can even be connected physically.

4. Intermediate Filaments

34Intermediate filament-forming proteins are a large protein class, encoded by at least 70 genes, organizing filaments with a diameter of 10 nm. Intermediate filaments are grouped in 5 classes according to their structure and sequence homology. Thereby, the first four classes represent cytoplasmic intermediate filaments, while type V are nuclear filaments, so called lamins (lamin A/C, B1, B2). Type I and II are acidic and basic keratins, forming heteropolymers consisting of a mixture of the 54 different type I and II keratins, expressed in dependence of cell type and differentiation status. In contrast, type III intermediate filaments are homopolymers of vimentin, desmin, peripherin, or glial fibrillary acidic protein (GFAP). Vimentin is mainly expressed in fibroblasts, endothelial cells, astrocytes; peripherin in neurons of the peripheral nervous system and desmin in muscle cells and GFAP mainly in astrocytes. Type IV intermediate filaments contain three neurofilament heteropolymers (NF-L/M/H), internexin, synemin, and nestin, mainly expressed in the cells of the nervous system. Nestin and synemin cannot form filaments on their own, but only in conjunction with other intermediate filament proteins. Two further intermediate filaments, called filensin and phakinin, cannot be grouped into the mentioned five types. They are expressed in the lens epithelium, forming heteropolymers. All cytoplasmic intermediate filaments have a similar monomer structure, consisting of a central α-helix with a non-helical structure at both of its ends. Two monomers spiral around each other, forming a so-called “coiled-coil” dimer and, subsequently, these dimers form unpolarized tetramers via antiparallel association and 8 tetramers form a cylindrical unit-filament. The unit filaments aggregate further with other unit filaments at the time scale of minutes to form intermediate filaments (Figure 5). After aggregation, the filaments undergo a compaction step during which the filament diameter shrinks to its final size of approximately 10 nm. For nucleation and polymerization of intermediate filaments co-factors are not needed. Intermediate filaments show a constant, but slow, subunit exchange along the whole filament, occurring at a rate of approximately 1 per 200 tetramers per hour in vitro for vimentin.

35Inside the cell, cytoplasmic intermediate filaments form a dense meshwork that is mainly located in the perinuclear space, but also reaches the cortex. The form and structure of the network depends on the type of intermediate filament. While keratin forms bundles and fibers that form only weakly connected networks, vimentin and desmin form highly connected networks with small mesh size and lamins generate filaments and fibers. Near the cortex, intermediate filaments interact with focal adhesion sites, desmosomes, and hemidesmosomes, maintaining cell and tissue adhesion. Conversely, desmosomes and focal adhesions function as centers for de novo intermediate filament formation. Via their anchorage with the nuclear and plasma membrane, intermediate filaments form a scaffold for mitochondria, the Golgi apparatus, and further organelles and organize their location. Due to its network structure and its ability to anchor organelles, intermediate filaments are often considered to be mechanical buffers. This idea is supported by the single filament properties of intermediate filaments that can withstand deformations of up to 300% of their initial length without rupturing. Looking at the elastic properties of intermediate filaments, they can be considered as flexible polymers with a persistence length of less than 1 µm. Interestingly, intermediate filaments show a strongly increasing elastic modulus with increasing deformation (called strain hardening). Measurements of single cells and simulations could confirm an important role of intermediate filaments for the overall visco-elastic response of a cell.

36Despite its function as a “mechanical buffer” and “organelle anchors”, intermediate filaments are highly dynamic components of the cytoskeleton, with multiple functions, including roles in apoptosis, migration, adhesion, and interactions with other cytoskeletal components. For fulfilling these functions, intermediate filaments need to form a defined network capable of (in-)direct interaction with its targets. Organizing factors are, amongst others, post-translational modifications as, for example, phosphorylation and acetylation, regulating assembly, organization, and function of intermediate filaments. A further influencer of intermediate filament organization is the plakin family of proteins, connecting microtubules and actin to intermediate filaments. Additionally, plakins connect intermediate filaments in desmosome adhesions and cell-matrix hemidesmosome adhesions to the actin and microtubule cytoskeleton and the nucleus and intermediate filaments to one another. Some intermediate filaments can also orient themselves along the actin or microtubule cytoskeleton. Actin and microtubules thereby form the guiding structures where filaments are transported along, either by kinesin and dynein (microtubules) or myosin (actin), resulting in a mutually dependent organization of intermediate filaments and actin and/or microtubules. Interestingly, vimentin transport along microtubules can be inhibited by the actin meshwork, demonstrating a complex interaction between the cytoskeletal components. Intermediate filaments are not only associated with other cytoskeletal proteins and (hemi-)desmosomes, but in the case of vimentin, it also binds via plectin or integrin α2β1 to actin and/or focal adhesions and promotes their strengthening. A direct interaction between the vimentin tail domain and actin is also proposed. The interactions of intermediate filaments discussed here and their interactions with microtubules and actin are summarized in Figure 2 and Figure 3 and Table 3.

37Intermediate filaments also link the nucleus to the cytoplasmic cytoskeleton via the LINC (linker of nucleoskeleton and cytoskeleton) complex that is present at the nuclear membrane, binding to plectin and, therewith, to intermediate filaments. Consequently, a disruption of LINC function leads to a disturbed force transmission due to the weakened linkage of the nucleus and cytoskeleton. Similarly, depletion of nestin, vimentin, and GFAP in astrocytes leads to positional and rotational changes of the nucleus. A correlation was found between nuclear rigidity, chromatin organization, and vimentin levels, indicating a crucial role of cytoplasmic intermediate filaments as passive mechanotransducers to the nucleus to control gene expression. Similarly, GFAP mutations and changes in desmin organization alter gene expression.

38Further types of intermediate filaments are the nuclear lamins. While nuclear lamins near the nuclear periphery form a filament network the organization of lamins in the center of the nucleus is only partly understood. It was proposed that the nucleoplasmic lamins may form filaments, short fibrous structures, foci, or an unstructured “veil”. The presence of seemingly less dense structures of lamins in the nucleoplasm is also in agreement with the observation of their higher mobility. Interestingly, a lamin A knockdown inhibits the expression of actomyosin cytoskeletal related genes, as shown in mesenchymal stem cells. Similarly, increased or reduced expression of lamin A inhibits or favors migration through a porous 3d matrix. On a functional level, A-type lamins seem to impact mechanosensing and signaling and to contribute to nuclear stiffness.

5. Involvement of Cytoskeleton in Cell Motility and Focal Adhesions

39One highly important property of a cell is its ability to move, especially conceivable in the context of immune cells chasing pathogens, wound closure, or metastasis of tumor cells. For cells to move efficiently, a few universal steps are necessary, as follows: It needs to form protrusions that attach to its surroundings and, subsequently, a contraction and retraction of the rear is necessary. A characteristic of cell migration is the precise coordination of these events in space and time. If, for example, the maturation of the adhesions at the cell front is not completed, an increase in contraction leads to the rupture of the newly formed adhesions, abandoning productive movement.

40To achieve productive movement and the right timing of migration steps, cells have developed two distinct modes of migration, the amoeboid and mesenchymal type. Amoeboid cell migration is characteristic for rounded cells with low adhesion and high Rho-driven contractility, whereas mesenchymal migrating cells show strong adhesion and Rac1-induced protrusions, in line with the mutual negative regulation of Rac1 and RhoA.

5.1. Actin in Motile Processes

41Actin filaments are the main contributors to cell migration in terms of force generation at the cell front and contraction at the rear. As stated, a cell can, in principle, use two types of migration, amoeboid and mesenchymal. While mesenchymal motion is mostly achieved via the extension of the lamellipodium or filopodia, the amoeboid migration works via the extension of blebs.

42The lamellipodium is one of the main force generating cell structures, generating pushing forces of up to 35 nN in the extreme case of fish keratocytes. To generate these forces, actin is polymerized locally at the cell front via Arp2/3 and depolymerized at the back of the lamellipodium by ADF/cofilin. Interestingly, formins such as FMNL2 or FMNL3, seem to participate in lamellipodial extension independent of Arp2/3 complex incorporation and are, in some cell types, major sources of lamellipodial protrusion forces. The protein Arpin inhibits the activity of Arp2/3, leading to pause phases in lamellipodial extensions and less directed motion. The continuous (de-)polymerization of actin creates a treadmilling effect and consequently forces and retrograde flow of actin. A flow in the opposite direction of the retrograde flow is generated by stress fiber contraction, transporting actin to the cell front. To hold the lamellipodium in place and prevent retraction via e.g., actin cortex tension, the formation of new cell-ECM contacts is necessary. Generally speaking, during lamellipodium extension nascent adhesions, that mature into focal adhesions or disassemble, form. While Rac1 controls the formation of nascent adhesions, maturation is controlled by RhoA and myosin II induced contractility, making them anchorage sites for stress fibers that generate tension and, thus, also control the composition of foal adhesions. The exact actin nucleation mechanism in focal adhesions is only partly understood, but formins, such as FHOD1 or mDia1, are supposed to play a role. The formation of nascent adhesions, in contrast, is thought to depend on Arp2/3 activity in the lamellipodium, due to its interaction with vinculin and focal adhesion kinases (FAK). Nevertheless, the lamellipodium is not essential for migration as several cell types, including fibroblasts and melanoblasts, can migrate without Rac or Arp2/3, but significantly slower. In the absence of the lamellipodium, these cells migrate via filopodia or other, probably formin dependent, pseudo-pods. For productive movement it is necessary to spatially restrict actin polymerization to one zone. Thus, it is assumed that Rac is only locally active. A mechanism to locally activate Rac1 is believed to function via Cdc42 induced pathways and microtubule capture at the leading edge and the subsequent local RacGEF and vesicle supply (see also next chapter). A further possible mechanism involves the Rho/ROCK pathway and actomyosin contractility to inhibit lamellipodium formation in multiple cell regions. This idea is supported by the occurrence of multiple or larger lamellipodia after inhibition of Rho or ROCK activity, giving a strong hint to the importance of a finely tuned dynamic equilibrium between contractile and expansive forces. Although RhoA/ROCK is active at the cell front, a too high activity impairs lamellipodium based migration via retraction of the lamellipodium.

43A second model, describing the generation of protrusions necessary for migration, is so-called blebbing. This mode of migration is observed in e.g., amoebae, tumor cells, neutrophils, or primordial germ cells or more generally in none- or weakly adherent cells, cells moving in a three dimensional matrix, or in confined environments. As previously described, blebs are initially actin free structures that arise by hydrostatic pressure causing a detachment of the actin-cortex from the membrane, thus extending the cell membrane. Hydrostatic pressure is generated mostly by actin cortex contraction. Local RhoA activation inducing myosin activation leads to increased contractility. The so generated hydrostatic pressure causes bleb formation through hydrostatic flow. Similarly to mesenchymal motion, cells using blebbing for migration need to “attach” the newly formed bleb to the surroundings and “detach” the cell rear, according to most models. It seems that adhesion of cells using blebs for migration is very low, implying that strong cell adhesions may actually impede bleb-based motion. Consequently, both a low adhesion and high cortex contractility favor the amoeboid motion type. One form of attachment of blebbing cells to their surroundings is via “chimneying”, which works via forces perpendicular to the direction of motion and, consequently, is independent of specific adhesion molecules. A different model proposed that forces are transmitted via cell-substrate intercalations. If blebs extend and form protrusions at the side of the cell into gaps of the substratum, then contractility of the reestablished cortex can than generate a net force to pull the cell body. Despite these models, other forms of bleb based propulsion have been suggested, like a flow friction driven or a “swimming in low Reynolds numbers” model. As bleb initiation and growth is mainly governed by myosin contractility and the actin cortex-membrane linkage, it is not surprising that the level of the actin-membrane cross-linker ezrin is increased at the cells rear and reduced at its front in carcinoma cells using blebbing for migration. Similarly, increasing the level of other ERM proteins impedes bleb formation and bleb induced migration, while reducing ERM protein levels has the opposite effect. A further factor that critically limits bleb extension and, thus, migratory properties is the cell membrane, that can usually stretch only about 4% before rupturing. As blebs do not normally contain endosomes, it is suggested that bleb expansion is allowed by the local unfolding of the membrane. Furthermore, bleb expansion is faster than lamellipodial growth, can occur in arbitrary directions, and, because they do not contain the cortex, they can naturally adapt to three dimensional environments. Therefore, blebs might be of high importance in complex three dimensional (in vivo) environments, where lamellipodial extension is seriously impeded.

44For efficient migration the cell rear needs to contract as well. To actively contract, actin structures use myosin to slide anti-parallel actin fibers along each other, creating contractile forces if the filaments are anchored at e.g., focal adhesions. The best studied contractile structures anchored to the substrate are stress fibers. Typically, stress fibers are directly linked to focal adhesions (except for transversal arcs), connecting the cell via actin fibers to the ECM. Interestingly, the formed focal adhesions are stress dependent and inhibition of myosin II generated contractility decreases focal adhesion size, while external tension favors focal adhesion maturation. Additionally, the forces acting on focal adhesions can lead to conformational changes of mechanosensitive proteins contained in focal adhesions, like β-integrins or talin, allowing stress fibers to convert mechanical into chemical signals, influencing focal adhesion maturation and turnover. Therefore, dorsal stress fibers help the maturation of focal adhesions via tension at the leading edge and ventral stress fibers at the trailing edge. Thus, stress fibers are highly important for cell adhesion but their function during cell migration remains poorly understood, as they are absent from many fast migrating cells, like leukocytes and Dictyostelium discoideum amoeba, as well as from cells embedded in soft three dimensional matrices. Consequently, it was proposed that stress fibers are not necessary for migration. Under certain circumstances, they might have an inhibitory effect on migration because the turnover is comparably slow and contractile forces may impede cell motion. Thus, the significance of stress fibers may be linked to their role in deforming the ECM, stabilizing focal adhesions, and through the generation of tension for rear contraction inside the cell. For rear retraction it seems as if the contractile forces generated by ventral stress fibers are of importance for the disassembly of posterior adhesions and an inhibition of protrusions at the rear. As stress fiber contractility in non-motile cells is associated with strengthening of focal adhesions, it needs to be tightly regulated to achieve just the right amount for the detachment of posterior adhesions. Consequently, a too strong RhoA activation inhibits cell migration via increased contractility and inhibition of contractility via ROCK inhibition can even increase motility in some cell types under specific conditions. The idea of rear retraction via stress fibers is further promoted by an adhesion gradient with lower adhesiveness at the rear.

45Taken together, actin or, to be more precise, the lamellipodium, filopodia, and blebs are the main causes of force generation for cell motility and contractile structures like stress fibers or the actin cortex are drivers of rear contraction.

5.2. Microtubules in Motile Processes

46In contrast to actin, microtubules are mostly not associated with force generation during migration, but rather with cell polarization and focal adhesions. The role of microtubules can, in principle, be divided into three categories, as follows: Participation in cell motility via their own mechanics, via signal transduction, and as a transport structure.

47Microtubules are capable of bearing high external pressure and, thus, help to maintain the cells’ shape under physiological conditions. In migrating cells, the microtubule (+)-end points in the direction of the plasma membrane and microtubules reaching the leading edge grow, at least in epithelial cells, more persistent, associated with EBs. As discussed before EBs can recruit further +TIPs that promote microtubule stabilization, protrusion formation, and cell migration. Other +TIPs, such as Clip-170 or its binding partner Clasp, act as rescue factors and increase the time of microtubules near the cortex. Stabilization of growth can lead to a more persistent force transmission, even though the maximal pushing force decreases quadratically with length, due to buckling. Notably, the stabilization of microtubules does not only promote a more persistent microtubule growth, but also a steadier supply with material needed for migration, as these microtubules persist longer near the leading edge, being ideal tracks for material supply, in agreement with the preference of some kinesin motors for microtubules stabilized by acetylation and detyrosination. Consequently, the polymerization of microtubules can generate a force of a few piconewton, on the same order as the force generated by motor proteins. An in vitro study demonstrated that the generated forces can indeed deform membranes. For a direct involvement of microtubules in the force generation process, a significant amount of microtubules actually have to reach the cell front. In most cell types, only very few microtubules reach the lamellipodium and the generated force is insufficient to generate large scale protrusions. In contrast, in neurons and astrocytes, microtubules are capable of generating protrusions. In neurons, microtubules are also found unattached to centrosomes, forming bundles and generating large enough forces to participate in axon formation. The free bundles (−)-ends are stabilized by members of the CAMSAP/Patronin family. Notably, in axons these bundles point with their (+)-end away from the cell body, thus being capable of creating larger forces than single microtubules, sufficient to promote neurite outgrowth.

48Cell motion associated structures have a high material consumption and, therefore, a steady supply is necessary to allow a continuous movement. For this transport, microtubules and their associated motor proteins are of high importance because they can transport membrane components needed for membrane extension, signaling molecules, such as the small GTPases Rac and Cdc42, GEFs, and proteases, but also intermediate filaments and their precursors. Additionally, microtubules transport β-actin coding mRNA and Arp2/3 subunits to the cell front.

49A further indirect way for microtubules to influence cell motility is via the (de-)stabilization of cell-matrix adhesions or regulation of actin (de-)polymerization, as microtubules are associated with focal adhesions, their regulation, RhoGTPase activity, and, consequently, actomyosin contractility. This can easily be seen by experiments with microtubule destabilizing agents like nocodazole, causing cell protrusion defects due to reduced Rac1 induced actin polymerization and increased contractility because of Rho-myosin II signaling. This is supported by observations showing that microtubule growth can locally activate Rac1, favoring the generation of new focal adhesion sites and the lamellipodium. In neuronal cells, Rac might be activated by TIAM1 (T-cell lymphoma invasion and metastasis-inducing protein 1) interacting with microtubules via MAP1B. An additional involvement of the Rac activators TRIO (triple functional domain protein) and TIAM2 was suggested in microtubule induced protrusion formation. A RhoGEF possibly involved in microtubule dependent regulation of RhoGTPases is H1, which interacts with microtubules, is inactive when bound, and transitions into an active state when microtubules depolymerize.

50Furthermore, microtubules grow in the direction of existing focal adhesions at the cell front where they get entrapped and stabilized and accelerate the maturation of focal adhesion via the transport of integrins. Thereby, stress fibers seem to function as a guidance structure for microtubules, mediated by MACF1. A positive feedback loop is also possible where integrin stimulation could cause a favored delivery of cargo at the site of adhesion. A further possibility is an interaction of FAK or paxilin with APC that clusters at microtubule tips. An additional signaling mechanism for APC and +TIPs, like Clip170 and CLASPs, is via the Rac and Cdc42 effector IQGAP and the formin mDia, promoting actin nucleation at focal adhesions. Despite the adhesion favoring effect of microtubules, an opposing destabilizing effect was observed at the cell rear. Microtubules actively targeted mature focal adhesions at the rear of motile fibroblasts, accelerating focal adhesion turnover. A common model describes the phenomenon via the dynamic instability of microtubule filaments. By growing and targeting focal adhesions, microtubules exert a force on the adhesion site, depolymerize quickly afterwards, and repeat the process. This hypothesis is supported by an observation showing a strong correlation between the microtubule poking number and the dissociation of focal adhesions. However, how microtubules find and target focal adhesions is not yet fully understood. It has been observed that they grow along actin bundles towards adhesion sites, potentially cross-linking to actin via e.g., spectraplakins or others. This idea is supported by the APC dependent localization of the spectraplakin MACF1 at the cell cortex, close to focal adhesions. Additionally, in the absence of MACF1 peripheral microtubules are less well organized and adhesion turnover is inhibited. Still, other mechanisms, involving CLASPs or interactions with integrin-linked kinase (ILK), are also possible. Furthermore, the actin bundling protein fascin interacts with microtubules, promoting focal adhesion turnover via FAK. Other mechanisms of microtubule dependent focal adhesion turnover are via clathrin mediated endocytosis of integrins, NBR1-mediated autophagy, or via vesicles carrying matrix metalloproteinases severing integrin-ECM connections.

5.3. Intermediate Filament Involvement in Motile Processes

51It is well established that intermediate filaments crucially influence both cell-matrix adhesion and migration. Nevertheless, the precise mechanism of action of intermediate filaments is not fully elucidated. As for microtubules it seems as though intermediate filaments are not a direct part of the force generation mechanism necessary for movement, but rather a signaling platform and mechanical anchor inside the cytoplasm to transduce forces through the whole cell.

52For this review we will focus the discussion mainly on vimentin, as vimentin is one of the best investigated intermediate filaments. Notably, we will not discuss keratins, as they are mostly restricted to epithelial cells and keratinocytes and not present in cell of glial origin.

53In non-migrating cells intermediate filaments are mostly localized around the nucleus, extending into the periphery, while they elongate into the lamella, connecting to focal adhesions near the leading edge in migrating cells. In the lamellipodium intermediate filaments are found mostly in a non-filamentous state. Generally speaking, intermediate filament organization alters the current state of the two other cytoskeletal components, thus potentially modulating both cell adhesion and migration.

54A strong hint for intermediate filaments influencing cell migration comes from the observation that, for example, vimentin can interact with actin and neurofilaments with microtubules. Vimentin is indeed necessary for motility of fibroblasts and breast cancer cells, epithelial cell wound closure, and other migration related phenomena. Vimentin inhibition reduces motility in fibroblasts, astrocytes, and diverse cancer cells. Studies evaluating possible mechanisms found evidence for vimentin directly binding to APC. APC regulates vimentin organization in astrocytes to align vimentin along the microtubule network. Post-translational modifications, like detyronsination and acetylation of microtubules, also impact vimentin network organization. On the other hand vimentin also affects polarized microtubule organization, amongst others by forming a template for microtubules, guiding microtubule growth, and, thus, favor directed migration. The exact mechanism of this interaction is not yet fully understood but may be governed by APC, linking microtubules and vimentin or via vimentin phosphorylation. Conversely, the vimentin filament network is dependent on microtubules and its motor proteins. Microtubule disruption leads to vimentin relocalization around the nucleus. Furthermore, activation of Cdc42 during scratch-wound assay in astrocytes inhibits dynein mediated rearward transport of GFAP and vimentin containing filaments, promoting intermediate filament network extension in direction of the leading edge.

55Another motility associated structure partly regulated by vimentin are focal adhesions. Vimentin is associated with formation, maturation, size, and strength of focal adhesions. Vimentin regulates the Rac1 GEF VAV2 and its localization to focal adhesions to promote their stabilization via Rac1 induced FAK activation. Depletion of vimentin in fibroblasts causes a FAK dependent induction of RhoA and myosin activity to compensate for the loss of tension induced by vimentin depletion. Similarly, vimentin depletion leads to increased stress fiber assembly and myosin activity, via RhoA activation by activating RhoA GEF-H1, but without activating FAK in osteosarcoma cells. Additionally, a triple silencing of vimentin, GFAP, and nestin or of vimentin and GFAP in astrocytes demonstrated that these intermediate filaments help to maintain the polarization of leader cells in collective motion by controlling forces in monolayers. Silencing of each individual intermediate filament produced similar but less pronounced results. This effect was attributed to larger and more focal adhesions that were distributed less concentratedly at the cell front. It is supposed that those three intermediate filaments control focal adhesions and traction force in astrocytes via plectin to control vinculin recruitment. In the case of vimentin, an interaction with integrin β3 was also observed. A further hypothesis of how intermediate filaments control focal adhesions and traction forces is via the acto-myosin network, by redirecting forces and restraining actin retrograde flow, or by regulation of focal adhesions via microtubules. Another mechanism that might explain the interaction of vimentin with focal adhesions is via the RAF-1/RhoA signaling, activating ROCK and being able to phosphorylate vimentin, leading to a filament collapse and subsequent release of ROCK at the cells periphery. Thus, the presence of vimentin can influence the RhoA signaling and, consequently, the formation and stability of focal adhesions.

56In line with these findings, transverse arcs interact via plectin with vimentin to promote their retrograde flow and this coupling is necessary for the perinuclear organization of vimentin filaments. Consequently, myosin II driven contractility of transversal arcs was made responsible for retrograde vimentin movement. This is of special interest, as the local activation of Rac1 causes a local disassembly of vimentin at the lamellipodium forming side, also preventing polymerization of vimentin via the Rac1 and Cdc42 effector PAK. Conversely, local vimentin depolymerization causes the formation of the lamellipodium at the side of vimentin depletion. Consequently, vimentin co-regulates both contractile actomyosin bundles and protrusive lamellipodial actin, and thus cell polarization. Despite the regulation of actin dynamics in the lamellipodium and lamellum, vimentin seems to form a transport structure for the nucleus in a three dimensional environment (“nuclear piston”) in cooperation with actomyosin, via the creation of a pressure gradient.

57Taken together, vimentin seems to mainly contribute to focal adhesion maturation and stabilization via the modulation of RhoA by FAK or Rho GEFs by maintenance of directed migration in collective motion and by regulation of the microtubule organization. Additionally, it seems as if vimentin locally suppresses the formation of the lamellipodium, favoring asymmetrical protrusion formation and, thus, generation of a net force.

58There have also been studies of other intermediate filaments, but the understanding of their impact on cell migration and adhesion is limited. Here we will only give some non-exhaustive examples. Synemin is associated with migration in astrocytoma via interactions with the focal adhesion associated protein zyxin and regulation of actin dynamics. Additionally, synemin was shown to interact with the focal adhesion components talin, vinculin, plectin, and α-actinin. Nestin was demonstrated to regulate FAK, integrin α5β1 localization and VASP activity, controlling invasiveness of prostate cancer cells.

6. Cytoskeletal Alterations in Glioma

59Malignancies belong to the most common causes of death worldwide, with increasing tendency. Among them, brain tumors occur at a rate of 5–10 per 100,000 individuals. The most frequent type of brain malignancies are gliomas that can be differentiated in astrocytoma, oligodendroglioma, and oligoastrocytoma. Especially high grade glioma are characterized by a highly infiltrative growth behavior, leading to low patient survival times. The infiltration of adjacent regions of the brain is one of the most critical parts in patients’ survival, as it makes complete tumor resection almost impossible, consequently leading to frequent recurrences.

60As migration and, thus, infiltration of glioma cells is largely governed by reshaping the cytoskeleton, it is no surprise that the composition and organization of the cytoskeleton in glioma cells differs strongly from that of healthy brain cells. In general, the Rho GTPases Rac1, Cdc42, and RhoA play a pivotal role in cell migration and cytoskeletal organization. The same is true for glioma cells, showing an increased expression of the mutually inhibiting GTPases RhoA and Rac1. Additionally, invading glioma cells exhibit an increased Cdc42 and Rac1 activity, but a decreased activity of RhoA at the cell front, held responsible for the enhanced migration of leader cells. Similarly, cells at the invading front show a markedly increased FAK expression. Thereby, the activity of RhoA seems to cause ambivalent effects. On the one hand, it is strongly expressed at the cell front and necessary for migration and, on the other hand, a marked activation causes an inhibition of glioma migration, via an inhibition of Rac1, and increased contractility, via larger focal adhesions. Inhibition of the RhoA target ROCK led to increased migration in glioma cells via an activation of Rac1 and Rac1 inhibition was associated with reduced migration and invasion. Contrasting these observations, other studies found ROCK inhibition associates with the reduced migration of glioma cells. A possible explanation for this discrepancy might be due to biphasic effects of ROCK activity. While low ROCK activity inhibits motion via low contractility and adhesion, a too high activity may impede migration due to contraction of protrusions and large adhesion sites, in a similar manner as proposed for the relation between adhesion and migration speed. The mutual inhibition of RhoA and Rac1 is further complicated by Cdc42 being capable of activating Rac1 in glioma. Interestingly, the formin mDia1, which is activated by RhoA, plays a major role in glioma cell polarization. For mDia1 expression, an association was demonstrated with the microtubule dependent localization of APC and Cdc42 at the cell front, regulating cell polarization. The impact of RhoA and Rac1 on glioma migration is in good agreement with the observation of an increased FAK activation, negatively regulating RhoA and activating Rac1 in glioma, being associated with staging and a poorer prognosis. In contrast, downregulation of FAK is associated with reduced migration. The important role of RhoA points to myosin as a further important player in glioma migration. Indeed, myosin was found to be crucial for glioma cell migration in vivo, likely for rear retraction and nucleus deformation. The lamellipodium is another structure associated with migration of glioma cells. Even though its main organizing protein Arp2/3 is associated with glioma staging, its exact role in glioma migration is not yet clear. While Arp2/3 inhibition was reported to reduce lamellipodial size and migration in a scratch wound assay, another study recognized no such relationship on linear functionalized tracks. Interestingly, Arp2/3 might also be involved in the maintenance of the glioma stem cell character. When cortactin, an important cross-linker in the dendritic actin network with overexpression in glioma, is inhibited the size of the lamellipodium and scratch wound closure is reduced.

61Silencing of the actin bundling protein fascin was demonstrated to reduce migratory capabilities of glioma cells, their division rate, and to increase the sensitivity to cytotoxic lymphocytes. The drop in migration was attributed to a loss of filopodia after fascin-1 silencing.

62Glioma cells are also capable of generating blebs for cell protrusion. The switch from mesenchymal to amoeboid motion occurs under conditions of low adhesion, via e.g., integrin β1 blockade, p130Cas silencing, or high contractility via RhoA activation. This is of special interest as migration capabilities of these cells was largely unchanged in 3D, pointing to an escape mechanism for interventions targeting “classical” migration associated molecules, such as e.g., integrin β1 or, potentially, others associated with mesenchymal migration. Still, the exact role of amoeboid motion in glioma invasion remains largely unknown.

63Glioma cells may express the intermediate filaments vimentin, GFAP, nestin, synemin, and α-internexin. Studies evaluating the organization and expression of intermediate filaments in glioblastoma found the following three distinct subpopulations with: (1) High vimentin, GFAP, and synemin expression or (2) low vimentin, GFAP, and synemin expression or (3) high nestin but low vimentin, GFAP, and synemin expression. Whether this expression pattern is associated with the origin of these tumors or has a distinct impact on migratory behavior is not yet elucidated. These patterns might alternatively be related with the molecular sub-classes of glioma (proneural, proliferative, mesenchymal). As opposed to healthy glia cells, glioma generally possess increased amounts of the intermediate filaments vimentin, nestin, and GFAP. While the amount of vimentin and the glioma stem cell marker nestin showed a negative correlation with patient prognosis and/or glioma staging, an association of GFAP expression with staging is controversial. A recent meta-study reported about a missing correlation between the amount of GFAP and the grade of gliomas. Interestingly, GFAP serum levels were proposed as a marker to differentiate between primary brain tumors and brain metastasis. The α-internexin levels correlated negatively with staging, but only in oligodendroglioma. These findings correlate well with the infiltrative or migratory phenotype obtained by glioma cells, as vimentin and nestin are associated with more motile characteristics (see before), while a forced expression of GFAP was shown to inhibit glioma motility in vitro. Vimentin may additionally have a role in radiation induced migration of glioma cells. After non-lethal irradiation, the expression of vimentin was increased and associated with faster migration. Furthermore, a nestin down-regulation is associated with an increased adhesion to collagen, fibronectin, and laminin, leading to an inhibition of migration of glioma cells, while overexpression had the reverse effect. This might be of further interest, as nestin was especially found in cells at the invasive front of glioma. The underlying molecular mechanisms of these nestin induced effects remained unclear so far. It remains tempting to speculate that, in glioma, a similar mechanism as discussed before might be responsible for the increased migratory capacity, via an activation of FAK and regulation of integrin localization. The intermediate filament synemin seems to be an additional part of the motile machinery of astrocytoma cells. Synemin is co-localized with the actin cross-linker α-actinin at the cell front and down-regulation of synemin impaired astrocytoma motility, via reduced F-actin and α-actinin amounts. Nevertheless, precise mechanisms of synemin actions in glioma are missing.

64Intermediate filaments are not only differentially expressed in glioma, but β-III, β-IV, and γ-tubulin are also overexpressed in glioma. Special attention should be given to β-III tubulin, which is not expressed in glia cells under physiologic conditions, but is up-regulated in high-grade glioma. β-III tubulin might be responsible for the resistance of glioblastoma to the microtubule stabilizing agent taxol, as observed in carcinoma cells, but this concept is still under debate. Notably, taxol was suggested to trigger differentiation in some glioma cells and in one study associated with an increase in GFAP expression. The higher expression of γ-tubulin might be related to centrosome abnormalities, altered microtubule dynamics, and consequently, adhesion dynamics and cell polarization. Furthermore, the microtubule severing protein spastin and the destabilizing protein stathmin are both associated with increased motility and, in case of spastin, also with reduced proliferation. Spastin expression is increased in glioma and mainly located at the cell front and the mitotic spindle, implying an indirect role in both motility and cell division by destabilization of microtubules, inducing spindle formation defects during division and facilitating microtubule turnover at the front to adapt to changes of the microenvironment. For the microtubule associated protein DCX different, contradictory results were found. Few studies reported that DCX is preferentially expressed at the invasive front of glioma, indicating a pro-invasive role. Other authors demonstrated a very low DCX expression in glioma and that a forced expression causes apoptosis and inhibits invasion. Accordingly, the role of DCX in glioma needs to be critically evaluated. In contrast, the role of EB1 seems to be unambiguous, the reduced accumulation of EB1 at microtubule tips is associated with higher microtubule instability, a less migratory and motile phenotype of glioma cells, caused by a lower amount of microtubules reaching the leading edge. The microtubule stabilizing factor APC, despite regulating differentiation and cell cycle arrest, might also be part of the migratory machinery of glioma cells, as it was recently shown that vimentin, GFAP, and nestin organization, along microtubules in the glioblastoma cell line U138-MG, is critically dependent on APC. APC may, additionally, increase the vimentin polymerization rate and, thus, influence migration. The motor protein dynein is not differentially expressed in glioma cells in general, but migrating glioma cells express increased amounts of the protein. Dynein is, in part, responsible for microtubule dependent intracellular transport and it can be speculated that dynein is necessary to maintain cell polarization by transporting inhibitory signals away from the cell front. Similarly, kinesin-5 and microtubule-actin linker protein MACF1 are involved in increased migration and up-regulated similar to KiF2C and KiF14 in glioma cells. The exact mechanism of MACF1 action is not elucidated yet, but a contribution of Wnt-signaling was proposed. Further microtubule and actin-associated proteins are differentially expressed in glioma. A summary can be found in Table 4.

65For productive migration, glioma cells generate an ECM differing from the normal micro-milieu of the central nervous system. This includes an overexpression of tenascin-C at the invasive front, increasing motility of glioma cells via interactions with β1 or αvβ3 integrins, probably via an activation of FAK and inhibition of RhoA. The idea of such a mechanism is supported by the observed drop in glioma migration after inhibition of β1- or αv-integrin and increased motility after β1 overexpression. In line with the association of integrins with migration is the elevated α-actinin expression in glioma, which is associated with a worse prognosis and connects focal adhesions to the cytoskeleton. Despite tenascin-C other ECM components were also associated with a more migratory phenotype of glioma, such as laminin, fibronectin, and collagen, but myelin biomembranes are also efficient substrates for migration. For thin linear laminin coatings trying to imitate properties of the basal lamina, a favored invasion route of glioma, it could be demonstrated that polarization of collective migration of glioma cells was microtubule dependent. This migration mode could largely be inhibited by formin inhibition, but not by inhibition of Arp2/3 that led to an increased migration speed. This is in line with the general importance of formins for glioma migration, demonstrating that mDia antagonism/depletion and agonism/overexpression impaired migration and invasion. The observed effect was, thereby, likely mediated via increased F-actin assembly and the stabilization of microtubules, leading to defects in protrusion formation, cell-cycle arrest, and apoptosis. Hyaluronic acid/hyaluronan (HA), the main ECM component of the healthy brain, as well as their main receptors CD44 and RHAMM (receptor of hyaluronan-mediated motility) are also upregulated in glioma, but their exact role is not yet determined. In general, increased HA expression is associated with an increase in motility and migration of glioma, probably via a CD44 or RHAMM induced activation of Rho GTPases, such as Rac1 or Cdc42. Even so, the role of CD44 remains ambivalent in glioma cell migration. On the one hand low and high levels of CD44 are associated with a less migratory phenotype and on the other hand intermediate expression levels facilitate migration. This biphasic effect is probably related to the substrate adhesiveness.

7. Conclusions

66The actin and microtubule cytoskeleton, their interactions, and relation to cell migration are comparably well understood for healthy cells, but the impact of their interplay with intermediate filaments just started to gain attention. Consequently, the role of intermediate filaments during migration is less well understood and many questions, especially regarding the role of GFAP, vimentin, synemin, and nestin in glia cells, remain open. For malignant cells the situations worsen, due to altered expression and localization profiles of cytoskeleton associated proteins, leading to changes in the global organization of the cytoskeleton, the signaling, and turnover of structures. Additional attention should be paid to the heterogeneity of glioma, showing significant differences in expression profile and behavior between “tumor-core” and “invading” tumor cells, but also between glioma of different patients. In case of glioma, the impact of cytoskeletal variations is only sparsely investigated and will need a significantly increased amount of data to understand the migration of glioma in the central nervous system. This includes the effect of the observed overexpressions, like spastin, vimentin, nestin, actin cross-linkers, etc., on migration and the underlying mechanisms, but also how migration associated signaling is altered in glioma and whether this can be used for specifically targeting glioma migration. Furthermore, the function of stress fibers in glioma migration and whether blebs play a significant role during brain infiltration by glioma are also not yet addressed adequately. Current approaches targeting the cytoskeleton in glioma mainly focus on microtubules, but mostly not to alter migratory properties but to induce cell cycle arrest or apoptosis. Nevertheless, these approaches remained futile because of glioma heterogeneity, therapeutic side effects, or resistance mechanisms.

References 612

1.
Seyfried T.N., Huysentruyt L.C. On the origin of cancer metastasis. Crit. Rev. Oncog. 2013;18:43-73.
Not in our library10.1615/critrevoncog.v18.i1-2.40
2.
Rotty J.D., Bear J.E. Competition and collaboration between different actin assembly pathways allows for homeostatic control of the actin cytoskeleton. Bioarchitecture. 2014;5:27-34.
Not in our library10.1080/19490992.2015.1090670
3.
Wen Q., Janmey P.A. Polymer physics of the cytoskeleton. Curr. Opin. Solid State Mater. Sci. 2011;15:177-182.
Not in our library10.1016/j.cossms.2011.05.002
4.
Sept D., Xu J., Pollard T.D., McCammon J.A. Annealing accounts for the length of actin filaments formed by spontaneous polymerization. Biophys. J. 1999;77:2911-2919.
Not in our library10.1016/s0006-3495(99)77124-9
5.
Sept D., Elcock A.H., McCammon J.A. Computer simulations of actin polymerization can explain the barbed-pointed end asymmetry. J. Mol. Biol. 1999;294:1181-1189.
Not in our library10.1006/jmbi.1999.3332
6.
Pollard T.D. Actin and Actin-Binding Proteins. Cold Spring Harb. Perspect. Biol. 2016;8:1-17.
Not in our library10.1101/cshperspect.a018226
7.
Pollard T.D., Blanchoin L., Mullins R.D. Molecular Mechanisms Controlling Actin Filament Dynamics in Nonmuscle Cells. Annu. Rev. Biophys. Biomol. Struct. 2000;29:545-576.
Not in our library10.1146/annurev.biophys.29.1.545
8.
Kang F., Purich D.L., Frederick S., Southwick F.S. Profilin Promotes Barbed-end Actin Filament Assembly without Lowering the Critical Concentration. J. Biol. Chem. 1999;274:36963-36972.
Not in our library10.1074/jbc.274.52.36963
9.
Machesky L.M., Mullins R.D., Higgs H.N., Kaiser D.A., Blanchoin L., May R.C., Hall M.E., Pollard T.D. Scar, a WASp-related protein, activates nucleation of actin filaments by the Arp2/3 complex. Proc. Natl. Acad. Sci. USA. 1999;96:3739-3744.
Not in our library10.1073/pnas.96.7.3739
10.
Kovar D.R., Harris E.S., Mahaffy R., Higgs H.N., Pollard T.D. Control of the assembly of ATP- and ADP-actin by formins and profilin. Cell. 2006;124:423-435.
Not in our library10.1016/j.cell.2005.11.038
11.
Romero S., Le Clainche C., Didry D., Egile C., Pantaloni D., Carlier M.F. Formin is a processive motor that requires profilin to accelerate actin assembly and associated ATP hydrolysis. Cell. 2004;119:419-429.
Not in our library10.1016/j.cell.2004.09.039
12.
Swaney K.F., Li R. Function and regulation of the Arp2/3 complex during cell migration in diverse environments. Curr. Opin. Cell Biol. 2016;42:63-72.
Not in our library10.1016/j.ceb.2016.04.005
13.
Svitkina T.M., Borisy G.G. Arp2/3 complex and actin depolymerizing factor/cofilin in dendritic organization and treadmilling of actin filament array in lamellipodia. J. Cell Biol. 1999;145:1009-1026.
Not in our library10.1083/jcb.145.5.1009
14.
Achard V., Martiel J.L., Michelot A., Guérin C., Reymann A.C., Blanchoin L., Boujemaa-Paterski R. A “Primer”-Based Mechanism Underlies Branched Actin Filament Network Formation and Motility. Curr. Biol. 2010;20:423-428.
Not in our library10.1016/j.cub.2009.12.056
15.
Molinie N., Gautreau A. The Arp2/3 Regulatory System and Its Deregulation in Cancer. Physiol. Rev. 2018;98:215-238.
Not in our library10.1152/physrev.00006.2017
16.
Akin O., Mullins R.D. Capping Protein Increases the Rate of Actin-Based Motility by Promoting Filament Nucleation by the Arp2/3 Complex. Cell. 2008;133:841-851.
Not in our library10.1016/j.cell.2008.04.011
17.
Kawska A., Carvalho K., Manzi J., Boujemaa-Paterski R., Blanchoin L., Martiel J.-L., Sykes C. How actin network dynamics control the onset of actin-based motility. Proc. Natl. Acad. Sci. USA. 2012;109:14440-14445.
Not in our library10.1073/pnas.1117096109
18.
Vignjevic D., Yarar D., Welch M.D., Peloquin J., Svitkina T., Borisy G.G. Formation of filopodia-like bundles in vitro from a dendritic network. J. Cell Biol. 2003;160:951-962.
Not in our library10.1083/jcb.200208059
19.
Bendix P.M., Koenderink G.H., Cuvelier D., Dogic Z., Koeleman B.N., Brieher W.M., Field C.M., Mahadevan L., Weitz D.A. A quantitative analysis of contractility in active cytoskeletal protein networks. Biophys. J. 2008;94:3126-3136.
Not in our library10.1529/biophysj.107.117960
20.
Koenderink G.H., Dogic Z., Nakamura F., Bendix P.M., MacKintosh F.C., Hartwig J.H., Stossel T.P., Weitz D.A. An active biopolymer network controlled by molecular motors. Proc. Natl. Acad. Sci. USA. 2009;106:15192-15197.
Not in our library10.1073/pnas.0903974106
21.
Finer J.T., Simmons R.M., Spudich J.A. Single myosin molecule mechanics: Piconewton forces and nanometre steps. Nature. 1994;368:113-119.
Not in our library10.1038/368113a0
22.
Mahajan R.K., Pardee J.D. Assembly mechanism of Dictyostelium myosin II: Regulation by K+, Mg2+, and actin filaments. Biochemistry. 1996;35:15504-15514.
Not in our library10.1021/bi9618981
23.
Matsumura F. Regulation of myosin II during cytokinesis in higher eukaryotes. Trends Cell Biol. 2005;15:371-377.
Not in our library10.1016/j.tcb.2005.05.004
24.
Jung H.S., Komatsu S., Ikebe M., Craig R. Head–Head and Head–Tail Interaction: A General Mechanism for Switching Off Myosin II Activity in Cells. Mol. Biol. Cell. 2008;19:3234-3242.
Not in our library10.1091/mbc.e08-02-0206
25.
Craig R., Smith R., Kendrick-Jones J. Light-chain phosphorylation controls the conformation of vertebrate non-muscle and smooth muscle myosin molecules. Nature. 1983;302.
Not in our library10.1038/302436a0
26.
Yumura S., Yoshida M., Betapudi V., Licate L.S., Iwadate Y., Nagasaki A., Uyeda T.Q.P., Egelhoff T.T. Multiple Myosin II Heavy Chain Kinases: Roles in Filament Assembly Control and Proper Cytokinesis in Dictyostelium. Mol. Biol. Cell. 2005;16:4256-4266.
Not in our library10.1091/mbc.e05-03-0219
27.
Murakami N., Kotula L., Hwang Y.W. Two distinct mechanisms for regulation of nonmuscle myosin assembly via the heavy chain: Phosphorylation for MIIB and Mts 1 binding for MIIA. Biochemistry. 2000;39:11441-11451.
Not in our library10.1021/bi000347e
28.
Murakami N., Chauhan V.P.S., Elzinga M. Two nonmuscle myosin II heavy chain isoforms expressed in rabbit brains: Filament forming properties, the effects of phosphorylation by protein kinase C and casein kinase II, and location of the phosphorylation sites. Biochemistry. 1998;37:1989-2003.
Not in our library10.1021/bi971959a
29.
Rosenberg M., Ravid S. Protein Kinase C Regulates Myosin IIB Phosphorylation, Cellular Localization, and Filament Assembly. Mol. Biol. Cell. 2006;17:1364-1374.
Not in our library10.1091/mbc.e05-07-0597
30.
Fletcher D.A., Mullins R.D. Cell mechanics and the cytoskeleton. Nature. 2010;463:485-492.
Not in our library10.1038/nature08908
31.
Köhler S., Schaller V., Bausch A.R. Collective dynamics of active cytoskeletal networks. PLoS ONE. 2011;6.
Not in our library10.1371/journal.pone.0023798
32.
Wagner B., Tharmann R., Haase I., Fischer M., Bausch A.R. Cytoskeletal polymer networks: The molecular structure of cross-linkers determines macroscopic properties. Proc. Natl. Acad. Sci. USA. 2006;103:13974-13978.
Not in our library10.1073/pnas.0510190103
33.
Falzone T.T., Lenz M., Kovar D.R., Gardel M.L. Assembly Kinetics Determine the Architecture of α-actinin Crosslinked F-actin Networks. Nat. Commun. 2013;3.
Not in our library10.1126/scisignal.274pe36.insulin
34.
Huang S. Arabidopsis VILLIN1 Generates Actin Filament Cables That Are Resistant to Depolymerization. Plant Cell Online. 2005;17:486-501.
Not in our library10.1105/tpc.104.028555
35.
Kasza K.E., Broedersz C.P., Koenderink G.H., Lin Y.C., Messner W., Millman E.A., Nakamura F., Stossel T.P., MacKintosh F.C., Weitz D.A. Actin filament length tunes elasticity of flexibly cross-linked actin networks. Biophys. J. 2010;99:1091-1100.
Not in our library10.1016/j.bpj.2010.06.025
36.
Klein M.G., Shi W., Ramagopal U., Tseng Y., Wirtz D., Kovar D.R., Staiger C.J., Almo S.C. Structure of the actin crosslinking core of fimbrin. Structure. 2004;12:999-1013.
Not in our library10.1016/j.str.2004.04.010
37.
Stossel T.P., Condeelis J., Cooley L., Hartwig J.H., Noegel A., Schleicher M., Shapiro S.S. Filamins as integrattors of cell mechanics and signalling. Nat. Rev. Mol. Cell Biol. 2001;2:138-145.
Not in our library10.1038/35052082
38.
Xu J., Schwarz W.H., Käs J.A., Stossel T.P., Janmey P.A., Pollard T.D. Mechanical properties of actin filament networks depend on preparation, polymerization conditions, and storage of actin monomers. Biophys. J. 1998;74:2731-2740.
Not in our library10.1016/s0006-3495(98)77979-2
39.
Bartles J.R. Parallel actin bundles and their multiple actin-bundling proteins. Curr. Opin. Cell Biol. 2000;12:72-78.
Not in our library10.1016/s0955-0674(99)00059-9
40.
Ferrer J.M., Lee H., Chen J., Pelz B., Nakamura F., Kamm R.D., Lang M.J. Measuring molecular rupture forces between single actin filaments and actin-binding proteins. Proc. Natl. Acad. Sci. USA. 2008;105:9221-9226.
Not in our library10.1073/pnas.0706124105
41.
Revenu C., Athman R., Robine S., Louvard D. The co-workers of actin filaments: From cell structures to signals. Nat. Rev. Mol. Cell Biol. 2004;5:635-646.
Not in our library10.1038/nrm1437
42.
Reymann A.C., Martiel J.L., Cambier T., Blanchoin L., Boujemaa-Paterski R., Théry M. Nucleation geometry governs ordered actin networks structures. Nat. Mater. 2010;9:827-832.
Not in our library10.1038/nmat2855
43.
Chesarone M.A., Dupage A.G., Goode B.L. Unleashing formins to remodel the actin and microtubule cytoskeletons. Nat. Rev. Mol. Cell Biol. 2010;11:62-74.
Not in our library10.1038/nrm2816
44.
Chen Q., Nag S., Pollard T.D. Formins filter modified actin subunits during processive elongation. J. Struct. Biol. 2012;177:32-39.
Not in our library10.1016/j.jsb.2011.10.005
45.
Goode B.L., Eck M.J. Mechanism and Function of Formins in the Control of Actin Assembly. Annu. Rev. Biochem. 2007;76:593-627.
Not in our library10.1146/annurev.biochem.75.103004.142647
46.
Michelot A., Derivery E., Paterski-Boujemaa R., Guérin C., Huang S., Parcy F., Staiger C.J., Blanchoin L. A Novel Mechanism for the Formation of Actin-Filament Bundles by a Nonprocessive Formin. Curr. Biol. 2006;16:1924-1930.
Not in our library10.1016/j.cub.2006.07.054
47.
Esue O., Harris E.S., Higgs H.N., Wirtz D. The Filamentous Actin Cross-Linking/Bundling Activity of Mammalian Formins. J. Mol. Biol. 2008;384:324-334.
Not in our library10.1016/j.jmb.2008.09.043
48.
Edwards M., Zwolak A., Schafer D.A., Sept D., Dominguez R., Cooper J.A. Capping protein regulators fine-tune actin assembly dynamics. Nat. Rev. Mol. Cell Biol. 2014;15:677-689.
Not in our library10.1038/nrm3869
49.
Laporte D., Ojkic N., Vavylonis D., Wu J.-Q. a-Actinin and fimbrin cooperate with myosin II to organize actomyosin bundles during contractile-ring assembly. Mol. Biol. Cell. 2012;23:3094-3110.
Not in our library10.1091/mbc.e12-02-0123
50.
Köhler S., Bausch A.R. Contraction mechanisms in composite active actin networks. PLoS ONE. 2012;7:1-8.
Not in our library10.1371/journal.pone.0039869
51.
Murrell M., Thoresen T., Gardel M. Reconstitution of contractile actomyosin arrays. Methods Enzymol. 2014;540:265-282.
Not in our library10.1016/b978-0-12-397924-7.00015-7
52.
Haviv L., Gillo D., Backouche F., Bernheim-Groswasser A. A Cytoskeletal Demolition Worker: Myosin II Acts as an Actin Depolymerization Agent. J. Mol. Biol. 2008;375:325-330.
Not in our library10.1016/j.jmb.2007.09.066
53.
Myosin motors fragment and compact membrane-bound actin filaments · 2013
No notice
54.
Reymann A.C., Boujemaa-Paterski R., Martiel J.L., Guérin C., Cao W., Chin H.F., De La Cruz E.M., Théry M., Blanchoin L. Actin network architecture can determine myosin motor activity. Science. 2012;336:1310-1314.
Not in our library10.1126/science.1221708
55.
Ingerman E., Hsiao J.Y., Mullins R.D. Arp2/3 complex ATP hydrolysis promotes lamellipodial actin network disassembly but is dispensable for assembly. J. Cell Biol. 2013;200:619-633.
Not in our library10.1083/jcb.201211069
56.
Reymann A.-C., Suarez C., Guerin C., Martiel J.-L., Staiger C.J., Blanchoin L., Boujemaa-Paterski R. Turnover of branched actin filament networks by stochastic fragmentation with ADF/cofilin. Mol. Biol. Cell. 2011;22:2541-2550.
Not in our library10.1091/mbc.e11-01-0052
57.
Andrianantoandro E., Pollard T.D. Mechanism of Actin Filament Turnover by Severing and Nucleation at Different Concentrations of ADF/Cofilin. Mol. Cell. 2006;24:13-23.
Not in our library10.1016/j.molcel.2006.08.006
58.
Cofilin-induced unidirectional cooperative conformational changes in actin filaments revealed by high-speed atomic force microscopy · 2015
No notice
59.
Elam W.A., Kang H., De La Cruz E.M. Biophysics of actin filament severing by cofilin. FEBS Lett. 2013;587:1215-1219.
Not in our library10.1016/j.febslet.2013.01.062
60.
Mccullough B.R., Blanchoin L., Martiel J., La E.M. De Cofilin Increases the Bending Flexibility of Actin Filaments: Implications for Severing and Cell Mechanics. J. Mol. Biol. 2009;381:550-558.
Not in our library10.1016/j.jmb.2008.05.055
61.
Cao W., Goodarzi J.P., De La Cruz E.M. Energetics and Kinetics of Cooperative Cofilin-Actin Filament Interactions. J. Mol. Biol. 2006;361:257-267.
Not in our library10.1016/j.jmb.2006.06.019
62.
Blanchoin L., Pollard T.D. Mechanism of interaction of Acanthamoeba actophorin (ADF/Cofilin) with actin filaments. J. Biol. Chem. 1999;274:15538-15546.
Not in our library10.1074/jbc.274.22.15538
63.
Kueh H.Y., Brieher W.M., Mitchison T.J. Quantitative analysis of actin turnover in listeria comet tails: Evidence for catastrophic filament turnover. Biophys. J. 2010;99:2153-2162.
Not in our library10.1016/j.bpj.2010.07.038
64.
Hao Y.K., Charras G.T., Mitchison T.J., Brieher W.M. Actin disassembly by cofilin, coronin, and Aip1 occurs in bursts and is inhibited by barbed-end cappers. J. Cell Biol. 2008;182:341-353.
Not in our library10.1083/jcb.200801027
65.
Suarez C., Roland J., Boujemaa-Paterski R., Kang H., McCullough B.R., Reymann A.C., Guérin C., Martiel J.L., De La Cruz E.M., Blanchoin L. Cofilin tunes the nucleotide state of actin filaments and severs at bare and decorated segment boundaries. Curr. Biol. 2011;21:862-868.
Not in our library10.1016/j.cub.2011.03.064
66.
Breitsprecher D., Koestler S.A., Chizhov I., Nemethova M., Mueller J., Goode B.L., Small J.V., Rottner K., Faix J. Cofilin cooperates with fascin to disassemble filopodial actin filaments. J. Cell Sci. 2011;124:3305-3318.
Not in our library10.1242/jcs.086934
67.
Schmoller K.M., Semmrich C., Bausch A.R. Slow down of actin depolymerization by cross-linking molecules. J. Struct. Biol. 2011;173:350-357.
Not in our library10.1016/j.jsb.2010.09.003
68.
Hayakawa K., Tatsumi H., Sokabe M. Actin filaments function as a tension sensor by tension-dependent binding of cofilin to the filament. J. Cell Biol. 2011;195:721-727.
Not in our library10.1083/jcb.201102039
69.
Chan C., Beltzner C.C., Pollard T.D. Cofilin Dissociates Arp2/3 Complex and Branches from Actin Filaments. Curr. Biol. 2009;19:537-545.
Not in our library10.1016/j.cub.2009.02.060
70.
Abercrombie M., Heaysman J.E.M., Pegrum S.M. Locomotion of fibroblasts in culture. V. Surface marking with concanavalin A. Exp. Cell Res. 1972;73:536-539.
Not in our library10.1016/0014-4827(72)90090-0
71.
Abercrombie M., Heaysman J.E.M., Pegrum S.M. The locomotion of fibroblasts in culture. IV. Electron microscopy of the leading lamella. Exp. Cell Res. 1971;67:359-367.
Not in our library10.1016/0014-4827(71)90420-4
72.
Lauffenburger D.A., Horwitz A.F. Cell migration: A physically integrated molecular process. Cell. 1996;84:359-369.
Not in our library10.1016/s0092-8674(00)81280-5
73.
Mitchison T.J., Cramer L.P. Actin-based cell motility and cell locomotion. Cell. 1996;84:371-379.
Not in our library10.1016/s0092-8674(00)81281-7
74.
Xu K., Babcock H.P., Zhuang X. Dual-objective STORM reveals three-dimensional filament organization in the actin cytoskeleton. Nat. Methods. 2012;9:185-188.
Not in our library10.1038/nmeth.1841
75.
Ydenberg C.A., Padrick S.B., Sweeney M.O., Gandhi M., Sokolova O., Goode B.L. GMF severs actin-Arp2/3 complex branch junctions by a cofilin- like mechanism. Curr. Biol. 2013;23:1037-1045.
Not in our library10.1016/j.cub.2013.04.058
76.
Vicente-Manzanares M., Ma X., Adelstein R.S., Horwitz A.R. Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat. Rev. Mol. Cell Biol. 2009;10:778-790.
Not in our library10.1038/nrm2786
77.
Vallotton P., Gupton S.L., Waterman-Storer C.M., Danuser G. Simultaneous mapping of filamentous actin flow and turnover in migrating cells by quantitative fluorescent speckle microscopy. Proc. Natl. Acad. Sci. USA. 2004;101:9660-9665.
Not in our library10.1073/pnas.0300552101
78.
Wilson C.A., Tsuchida M.A., Allen G.M., Barnhart E.L., Applegate K.T., Yam P.T., Ji L., Keren K., Danuser G., Theriot J.A. Myosin II contributes to cell-scale actin network treadmilling via network disassembly. Nature. 2010;465:373-377.
Not in our library10.1038/nature08994
79.
Prass M., Jacobson K., Mogilner A., Radmacher M. Direct measurement of the lamellipodial protrusive force in a migrating cell. J. Cell Biol. 2006;174:767-772.
Not in our library10.1083/jcb.200601159
80.
Krause M., Gautreau A. Steering cell migration: Lamellipodium dynamics and the regulation of directional persistence. Nat. Rev. Mol. Cell Biol. 2014;15:577-590.
Not in our library10.1038/nrm3861
81.
Tang H., Li A., Bi J., Veltman D.M., Zech T., Spence H.J., Yu X., Timpson P., Insall R.H., Frame M.C., et al. Loss of scar/WAVE complex promotes N-WASP- and FAK-dependent invasion. Curr. Biol. 2013;23:107-117.
Not in our library10.1016/j.cub.2012.11.059
82.
Petrie R.J., Gavara N., Chadwick R.S., Yamada K.M. Nonpolarized signaling reveals two distinct modes of 3D cell migration. J. Cell Biol. 2012;197:439-455.
Not in our library10.1083/jcb.201201124
83.
Breitsprecher D., Kiesewetter A.K., Linkner J., Vinzenz M., Stradal T.E.B., Small J.V., Curth U., Dickinson R.B., Faix J. Molecular mechanism of Ena/VASP-mediated actin-filament elongation. EMBO J. 2011;30:456-467.
Not in our library10.1038/emboj.2010.348
84.
Hansen S.D., Mullins R.D. VASP is a processive actin polymerase that requires monomeric actin for barbed end association. J. Cell Biol. 2010;191:571-584.
Not in our library10.1083/jcb.201003014
85.
Mejillano M.R., Kojima S.I., Applewhite D.A., Gertler F.B., Svitkina T.M., Borisy G.G. Lamellipodial versus filopodial mode of the actin nanomachinery: Pivotal role of the filament barbed end. Cell. 2004;118:363-373.
Not in our library10.1016/j.cell.2004.07.019
86.
Small J.V., Herzog M., Anderson K. Actin filament organization in the fish keratocyte lamellipodium. J. Cell Biol. 1995;129:1275-1286.
Not in our library10.1083/jcb.129.5.1275
87.
Helgeson L.A., Nolen B.J. Mechanism of synergistic activation of Arp2/3 complex by cortactin and N-WASP. Elife. 2013;2013:1-26.
Not in our library10.7554/elife.00884
88.
Dang I., Gorelik R., Sousa-Blin C., Derivery E., Guérin C., Linkner J., Nemethova M., Dumortier J.G., Giger F.A., Chipysheva T.A., et al. Inhibitory signalling to the Arp2/3 complex steers cell migration. Nature. 2013;503:281-284.
Not in our library10.1038/nature12611
89.
Veltman D. Actin dynamics: Cell migration takes a new turn with arpin. Curr. Biol. 2014;24:R31-R33.
Not in our library10.1016/j.cub.2013.11.022
90.
Houk A.R., Jilkine A., Mejean C.O., Boltyanskiy R., Dufresne E.R., Angenent S.B., Altschuler S.J., Wu L.F., Weiner O.D. Membrane tension maintains cell polarity by confining signals to the leading edge during neutrophil migration. Cell. 2012;148:175-188.
Not in our library10.1016/j.cell.2011.10.050
91.
Ofer N., Mogilner A., Keren K. Actin disassembly clock determines shape and speed of lamellipodial fragments. PNAS. 2011;108:20394-20399.
Not in our library10.1073/pnas.1105333108
92.
Svitkina T.M., Verkhovsky A.B., McQuade K.M., Borisy G.G. Analysis of the actin-myosin II system in fish epidermal keratocytes: Mechanism of cell body translocation. J. Cell Biol. 1997;139:397-415.
Not in our library10.1083/jcb.139.2.397
93.
Weichsel J., Urban E., Small J.V., Schwarz U.S. Reconstructing the orientation distribution of actin filaments in the lamellipodium of migrating keratocytes from electron microscopy tomography data. Cytom. Part A. 2012;81 A:496-507.
Not in our library10.1002/cyto.a.22050
94.
Faix J., Rottner K. The making of filopodia. Curr. Opin. Cell Biol. 2006;18:18-25.
Not in our library10.1016/j.ceb.2005.11.002
95.
Cojoc D., Difato F., Ferrari E., Shahapure R.B., Laishram J., Righi M., Di Fabrizio E.M., Torre V. Properties of the force exerted by filopodia and lamellipodia and the involvement of cytoskeletal components. PLoS ONE. 2007;2.
Not in our library10.1371/journal.pone.0001072
96.
Jacquemet G., Hamidi H., Ivaska J. Filopodia in cell adhesion, 3D migration and cancer cell invasion. Curr. Opin. Cell Biol. 2015;36:23-31.
Not in our library10.1016/j.ceb.2015.06.007
97.
Vignjevic D., Peloquin J., Borisy G.G. In vitro assembly of filopodia-like bundles. Methods Enzymol. 2006;406:727-739.
Not in our library10.1016/s0076-6879(06)06057-5
98.
Yang C., Svitkina T. Filopodia initiation: Focus on the Arp2/3 complex and formins. Cell Adhes. Migr. 2011;5:402-408.
Not in our library10.4161/cam.5.5.16971
99.
Block J., Breitsprecher D., Kühn S., Winterhoff M., Kage F., Geffers R., Duwe P., Rohn J.L., Baum B., Brakebusch C., et al. FMNL2 drives actin-based protrusion and migration downstream of Cdc42. Curr. Biol. 2012;22:1005-1012.
Not in our library10.1016/j.cub.2012.03.064
100.
Miki H., Sasaki T., Takai Y., Takenawa T. Induction of filopodium formation by a WASP-related actin-depolymerizing protein N-WASP. Nature. 1998;391:93-96.
Not in our library10.1038/34208
101.
Young L.E., Heimsath E.G., Higgs H.N. Cell type-dependent mechanisms for formin-mediated assembly of filopodia. Mol. Biol. Cell. 2015;26:4646-4659.
Not in our library10.1091/mbc.e15-09-0626
102.
Khurana S., George S.P. The role of actin bundling proteins in the assembly of filopodia in epithelial cells. Cell Adh. Migr. 2011;5:409-420.
Not in our library10.4161/cam.5.5.17644
103.
Chan C.E., Odde D.J. Traction dynamics of filopodia on compliant substrates. Science. 2008;322:1687-1691.
Not in our library10.1126/science.1163595
104.
Kress H., Stelzer E.H.K., Holzer D., Buss F., Griffiths G., Rohrbach A. Filopodia act as phagocytic tentacles and pull with discrete steps and a load-dependent velocity. Proc. Natl. Acad. Sci. USA. 2007;104:11633-11638.
Not in our library10.1073/pnas.0702449104
105.
Naumanen P., Lappalainen P., Hotulainen P. Mechanisms of actin stress fibre assembly. J. Microsc. 2008;231:446-454.
Not in our library10.1111/j.1365-2818.2008.02057.x
106.
Cramer L.P., Siebert M., Mitchison T.J. Identification of novel graded polarity actin filament bundles in locomoting heart fibroblasts: Implications for the generation of motile force. J. Cell Biol. 1997;136:1287-1305.
Not in our library10.1083/jcb.136.6.1287
107.
Sjöblom B., Salmazo A., Djinović-Carugo K. α-Actinin structure and regulation. Cell. Mol. Life Sci. 2008;65:2688-2701.
Not in our library10.1007/s00018-008-8080-8
108.
Koenderink G.H., Paluch E.K. Architecture shapes contractility in actomyosin networks. Curr. Opin. Cell Biol. 2018;50:79-85.
Not in our library10.1016/j.ceb.2018.01.015
109.
Pellegrin S., Mellor H. Actin stress fibres. J. Cell Sci. 2007;120:3491-3499.
Not in our library10.1242/jcs.018473
110.
Hotulainen P., Lappalainen P. Stress fibers are generated by two distinct actin assembly mechanisms in motile cells. J. Cell Biol. 2006;173:383-394.
Not in our library10.1083/jcb.200511093
111.
Tojkander S., Gateva G., Schevzov G., Hotulainen P., Naumanen P., Martin C., Gunning P.W., Lappalainen P. A molecular pathway for myosin II recruitment to stress fibers. Curr. Biol. 2011;21:539-550.
Not in our library10.1016/j.cub.2011.03.007
112.
Endlich N., Schordan E., Cohen C.D., Kretzler M., Lewko B., Welsch T., Kriz W., Otey C.A., Endlich K. Palladin is a dynamic actin-associated protein in podocytes. Kidney Int. 2009;75:214-226.
Not in our library10.1038/ki.2008.486
113.
Schmidt K., Nichols B.J. Functional interdependence between septin and actin cytoskeleton. BMC Cell Biol. 2004;5:1-13.
Not in our library10.1186/1471-2121-5-43
114.
Lazarides E., Burridge K. α-Actinin: Immunofluorescent localization of a muscle structural protein in nonmuscle cells. Cell. 1975;6:289-298.
Not in our library10.1016/0092-8674(75)90180-4
115.
Vallenius T., Makela T.P. Clik1: A novel kinase targeted to actin stress fibers by the CLP-36 PDZ-LIM protein. J. Cell Sci. 2002;115:2067-2073.
Not in our library10.1242/jcs.115.10.2067
116.
Vallenius T., Luukko K., Mäkelä T.P. CLP-36 PDZ-LIM Protein Associates with Nonmuscle alpha -Actinin-1 and alpha -Actinin-4. J. Biol. Chem. 2000;275:11100-11105.
Not in our library10.1074/jbc.275.15.11100
117.
Wang K., Ash J.F., Singer S.J. Filamin, a New High-Molecular-Weight Protein Found in Smooth Muscle and Nonmuscle Cells. Purification and Properties of Chicken Gizzard Filamin. Proc. Natl. Acad. Sci. USA. 1975;72:4483-4486.
Not in our library10.1073/pnas.72.11.4483
118.
Dixon R.D.S., Arneman D.K., Rachlin A.S., Sundaresan N.R., Costello M.J., Campbell S.L., Otey C.A. Palladin is an actin cross-linking protein that uses immunoglobulin-like domains to bind filamentous actin. J. Biol. Chem. 2008;283:6222-6231.
Not in our library10.1074/jbc.m707694200
119.
Elkhatib N., Neu M.B., Zensen C., Schmoller K.M., Louvard D., Bausch A.R., Betz T., Vignjevic D.M. Fascin plays a role in stress fiber organization and focal adhesion disassembly. Curr. Biol. 2014;24:1492-1499.
Not in our library10.1016/j.cub.2014.05.023
120.
Boukhelifa M., Moza M., Johansson T., Rachlin A., Parast M., Huttelmaier S., Roy P., Jockusch B.M., Carpen O., Karlsson R., et al. The proline-rich protein palladin is a binding partner for profilin. FEBS J. 2006;273:26-33.
Not in our library10.1111/j.1742-4658.2005.05036.x
121.
Boukhelifa M., Parast M.M., Bear J.E., Gertler F.B., Otey C.A. Palladin Is a Novel Binding Partner for Ena/VASP Family Members. Cell Motil. Cytoskeleton. 2004;58:17-29.
Not in our library10.1002/cm.10173
122.
Strasser P., Gimona M., Moessler H., Herzog M., Small J.V. Mammalian calponin: Identification and expression of genetic variants. FEBS J. 1993;330:13-18.
Not in our library10.1016/0014-5793(93)80909-e
123.
Weber K., Groeschel-Stewart U. Antibody to myosin: The specific visualization of myosin-containing filaments in nonmuscle cells. Proc. Natl. Acad. Sci. USA. 1974;71:4561-4564.
Not in our library10.1073/pnas.71.11.4561
124.
Yamashiro-Matsumura S., Matsumura F. Characterization of 83-kilodalton nonmuscle caldesmon from cultured rat cells: Stimulation of actin binding of nonmuscle tropomyosin and periodic localization along microfilaments like tropomyosin. J. Cell Biol. 1988;106:1973-1983.
Not in our library10.1083/jcb.106.6.1973
125.
Castellino F., Ono S., Matsumura F., Luini A. Essential role of caldesmon in the actin filament reorganization induced by glucocorticoids. J. Cell Biol. 1995;131:1223-1230.
Not in our library10.1083/jcb.131.5.1223
126.
Leung T., Chen X.Q., Manser E., Lim L. The p160 RhoA-binding kinase ROK alpha is a member of a kinase family and is involved in the reorganization of the cytoskeleton. Mol. Cell. Biol. 1996;16:5313-5327.
Not in our library10.1128/mcb.16.10.5313
127.
Watanabe N., Madaule P., Reid T., Ishizaki T., Watanabe G., Kakizuka A., Saito Y., Nakao K., Jockusch B.M., Narumiya S. p140mDia, a mammalian homolog of Drosophila diaphanous, is a target protein for Rho small GTPase and is a ligand for profilin. EMBO J. 1997;16:3044-3056.
Not in our library10.1093/emboj/16.11.3044
128.
Tominaga T., Sahai E., Chardin P., McCormick F., Courtneidge S.A., Alberts A.S. Diaphanous-related formins bridge Rho GTPase and Src tyrosine kinase signaling. Mol. Cell. 2000;5:13-25.
Not in our library10.1016/s1097-2765(00)80399-8
129.
Maekawa M., Ishizaki T., Boku S., Watanabe N., Fujita A., Iwamatsu A., Obinata T., Ohashi K., Mizuno K., Narumiya S. Signaling from Rho to the actin cytoskeleton through protein kinases ROCK and LIM-kinase. Science. 1999;285:895-898.
Not in our library10.1126/science.285.5429.895
130.
Chen Z., Borek D., Padrick S.B., Gomez T.S., Metlagel Z., Ismail A.M., Umetani J., Billadeau D.D., Otwinowski Z., Rosen M.K. Structure and control of the actin regulatory WAVE complex. Nature. 2010;468:533-538.
Not in our library10.1038/nature09623
131.
Lebensohn A.M., Kirschner M.W. Activation of the WAVE Complex by Coincident Signals Controls Actin Assembly. Mol. Cell. 2009;36:512-524.
Not in our library10.1016/j.molcel.2009.10.024
132.
Campellone K.G., Welch M.D. A nucleator arms race: Cellular control of actin assembly. Nat. Rev. Mol. Cell Biol. 2010;11:237-251.
Not in our library10.1038/nrm2867
133.
Peterson L.J., Rajfur Z., Maddox A.S., Freel C.D., Chen Y., Edlund M., Otey C., Burridge K. Simultaneous Stretching and Contraction of Stress Fibers In Vivo. Mol. Biol. Cell. 2004;15:3497-3508.
Not in our library10.1091/mbc.e03-09-0696
134.
Lu L., Oswald S.J., Ngu H., Yin F.C.P. Mechanical properties of actin stress fibers in living cells. Biophys. J. 2008;95:6060-6071.
Not in our library10.1529/biophysj.108.133462
135.
Deguchi S., Ohashi T., Sato M. Tensile properties of single stress fibers isolated from cultured vascular smooth muscle cells. J. Biomech. 2006;39:2603-2610.
Not in our library10.1016/j.jbiomech.2005.08.026
136.
Anderson T.W., Vaughan A.N., Cramer L.P. Retrograde Flow and Myosin II Activity within the Leading Cell Edge Deliver F-Actin to the Lamella to Seed the Formation of Graded Polarity Actomyosin II Filament Bundles. Mol. Biol. Cell. 2008;19:5006-5018.
Not in our library10.1091/mbc.e08-01-0034
137.
Langanger G., Moeremans M., Daneels G., Sobieszek A., De Brabander M., De Mey J. The molecular organisation of myosin in stress fibers of cultured cells. J. Cell Biol. 1986;102:200-209.
Not in our library10.1083/jcb.102.1.200
138.
Burnette D.T., Manley S., Sengupta P., Sougrat R., Davidson M.W., Kachar B., Lippincott-Schwartz J. A role for actin arcs in the leading-edge advance of migrating cells. Nat. Cell Biol. 2011;13:371-382.
Not in our library10.1038/ncb2205
139.
Hu S., Dasbiswas K., Guo Z., Tee Y.H., Thiagarajan V., Hersen P., Chew T.L., Safran S.A., Zaidel-Bar R., Bershadsky A.D. Long-range self-organization of cytoskeletal myosin II filament stacks. Nat. Cell Biol. 2017;19:133-141.
Not in our library10.1038/ncb3466
140.
Dasanayake N.L., Michalski P.J., Carlsson A.E. General mechanism of actomyosin contractility. Phys. Rev. Lett. 2011;107:1-4.
Not in our library10.1103/physrevlett.107.118101
141.
Nemethova M., Auinger S., Small J.V. Building the actin cytoskeleton: Filopodia contribute to the construction of contractile bundles in the lamella. J. Cell Biol. 2008;180:1233-1244.
Not in our library10.1083/jcb.200709134
142.
Generation of contractile actomyosin bundles depends on mechanosensitive actin filament assembly and disassembly · 2015
No notice
143.
Small J.V., Rottner K., Kaverina I., Anderson K.I. Assembling an actin cytoskeleton for cell attachment and movement. Biochim. Biophys. Acta Mol. Cell Res. 1998;1404:271-281.
Not in our library10.1016/s0167-4889(98)00080-9
144.
Chen W.-T. Mechanism of the trailing edge during fibroblast movement. J.Cell Biol. 1981;90:187-200.
Not in our library10.1083/jcb.90.1.187
145.
Khatau S.B., Hale C.M., Stewart-Hutchinson P.J., Patel M.S., Stewart C.L., Searson P.C., Hodzic D., Wirtz D. A perinuclear actin cap regulates nuclear shape. Proc. Natl. Acad. Sci. USA. 2009;106:19017-19022.
Not in our library10.1073/pnas.0908686106
146.
Burridge K., Guilluy C. Focal adhesions, stress fibers and mechanical tension. Exp. Cell Res. 2016;343:14-20.
Not in our library10.1016/j.yexcr.2015.10.029
147.
Gateva G., Tojkander S., Koho S., Carpen O., Lappalainen P. Palladin promotes assembly of non-contractile dorsal stress fibers through VASP recruitment. J. Cell Sci. 2014;127:1887-1898.
Not in our library10.1242/jcs.135780
148.
Kovac B., Teo J.L., Mäkelä T.P., Vallenius T. Assembly of non-contractile dorsal stress fibers requires α-actinin-1 and Rac1 in migrating and spreading cells. J. Cell Sci. 2013;126:263-273.
Not in our library10.1242/jcs.115063
149.
Shemesh T., Verkhovsky A.B., Svitkina T.M., Bershadsky A.D., Kozlov M.M. Role of focal adhesions and mechanical stresses in the formation and progression of the lamellum interface. Biophys. J. 2009;97:1254-1264.
Not in our library10.1016/j.bpj.2009.05.065
150.
Eghiaian F., Rigato A., Scheuring S. Structural, mechanical, and dynamical variability of the actin cortex in living cells. Biophys. J. 2015;108:1330-1340.
Not in our library10.1016/j.bpj.2015.01.016
151.
Clausen M.P., Colin-York H., Schneider F., Eggeling C., Fritzsche M. Dissecting the actin cortex density and membrane-cortex distance in living cells by super-resolution microscopy. J. Phys. D Appl. Phys. 2017;50.
Not in our library10.1088/1361-6463/aa52a1
152.
Hanakam F., Albrecht R., Eckerskorn C., Matzner M., Gerisch G. Myristoylated and non-myristoylated forms of the pH sensor protein hisactophilin II: Intracellular shuttling to plasma membrane and nucleus monitored in real time by a fusion with green fluorescent protein. EMBO J. 1996;15:2935-2943.
Not in our library10.1002/j.1460-2075.1996.tb00656.x
153.
Morone N., Usukura J., Kusumi A. Three-dimensional reconstruction of the membrane skelton at the plama membrane interface by electron tomography. J. Cell Biol. 2006;174:851-862.
Not in our library10.1083/jcb.200606007
154.
Biro M., Romeo Y., Kroschwald S., Bovellan M., Boden A., Tcherkezian J., Roux P.P., Charras G., Paluch E.K. Cell cortex composition and homeostasis resolved by integrating proteomics and quantitative imaging. Cytoskeleton. 2013;70:741-754.
Not in our library10.1002/cm.21142
155.
Cellular Control of Cortical Actin Nucleation · 2014
No notice
156.
Roh-johnson M., Shemer G., Higgins C.D., Mcclellan J.H., Werts A.D., Tulu U.S., Gao L., Betzig E., Kiehart D.P., Goldstein B. Triggering a cell shape change by exploiting preexisting actomyosin contractions. Science. 2012;335:1232-1235.
Not in our library10.1126/science.1217869
157.
Charras G.T., Hu C.K., Coughlin M., Mitchison T.J. Reassembly of contractile actin cortex in cell blebs. J. Cell Biol. 2006;175:477-490.
Not in our library10.1083/jcb.200602085
158.
McClatchey A.I., Fehon R.G. Merlin and the ERM proteins - regulators of receptor distribution and signaling at the cell cortex. Trends Cell Biol. 2009;19:198-206.
Not in our library10.1016/j.tcb.2009.02.006
159.
Carlier M., Laurent V., Santolini J., Melki R., Didry D., Xia G., Hong Y., Chua N., Pantaloni D. Actin depolymerizing factor (ADF/Cofilin) enhances the rate of filament turnover: Implication in actin-based motility. J. Cell Biol. 1997;136:1307-1322.
Not in our library10.1083/jcb.136.6.1307
160.
Chugh P., Clark A.G., Smith M.B., Cassani D.A.D., Dierkes K., Ragab A., Roux P.P., Charras G., Salbreux G., Paluch E.K. Actin cortex architecture regulates cell surface tension. Nat. Cell Biol. 2017;19:689-697.
Not in our library10.1038/ncb3525
161.
Howard J. Mechanics of Motor Proteins and the Cytoskeleton. 2005.
Not in our library
162.
Ananthakrishnan R., Guck J., Wottawah F., Schinkinger S., Lincoln B., Romeyke M., Moon T., Käs J. Quantifying the contribution of actin networks to the elastic strength of fibroblasts. J. Theor. Biol. 2006;242:502-516.
Not in our library10.1016/j.jtbi.2006.03.021
163.
Tee S.Y., Fu J., Chen C.S., Janmey P.A. Cell shape and substrate rigidity both regulate cell stiffness. Biophys. J. 2011;100:L25-L27.
Not in our library10.1016/j.bpj.2010.12.3744
164.
Guha M., Zhou M., Wang Y.L. Cortical actin turnover during cytokinesis requires myosin II. Curr. Biol. 2005;15:732-736.
Not in our library10.1016/j.cub.2005.03.042
165.
Murthy K., Wadsworth P. Myosin-II-dependent localization and dynamics of F-actin during cytokinesis. Curr. Biol. 2005;15:724-731.
Not in our library10.1016/j.cub.2005.02.055
166.
Bursac P., Lenormand G., Fabry B., Oliver M., Weitz D.A., Viasnoff V., Butler J.P., Fredberg J.J. Cytoskeletal remodelling and slow dynamics in the living cell. Nat. Mater. 2005;4:557-561.
Not in our library10.1038/nmat1404
167.
Gardel M.L., Nakamura F., Hartwig J.H., Crocker J.C., Stossel T.P., Weitz D.A. Prestressed F-actin networks cross-linked by hinged filamins replicate mechanical properties of cells. Proc. Natl. Acad. Sci. USA. 2006;103:1762-1767.
Not in our library10.1073/pnas.0504777103
168.
Lecuit T., Lenne P.F. Cell surface mechanics and the control of cell shape, tissue patterns and morphogenesis. Nat. Rev. Mol. Cell Biol. 2007;8:633-644.
Not in our library10.1038/nrm2222
169.
Charras G.T., Coughlin M., Mitchison T.J., Mahadevan L. Life and times of a cellular bleb. Biophys. J. 2008;94:1836-1853.
Not in our library10.1529/biophysj.107.113605
170.
Kasza K.E., Rowat A.C., Liu J., Angelini T.E., Brangwynne C.P., Koenderink G.H., Weitz D.A. The cell as a material. Curr. Opin. Cell Biol. 2007;19:101-107.
Not in our library10.1016/j.ceb.2006.12.002
171.
The role and regulation of blebs in cell migration · 2013
No notice
172.
Goudarzi M., Banisch T.U., Mobin M.B., Maghelli N., Tarbashevich K., Strate I., van den Berg J., Blaser H., Bandemer S., Paluch E., et al. Identification and Regulation of a Molecular Module for Bleb-Based Cell Motility. Dev. Cell. 2012;23:210-218.
Not in our library10.1016/j.devcel.2012.05.007
173.
Tinevez J.-Y., Schulze U., Salbreux G., Roensch J., Joanny J.-F., Paluch E. Role of cortical tension in bleb growth. Proc. Natl. Acad. Sci. USA. 2009;106:18581-18586.
Not in our library10.1073/pnas.0903353106
174.
Paluch E., Piel M., Prost J., Bornens M., Sykes C. Cortical actomyosin breakage triggers shape oscillations in cells and cell fragments. Biophys. J. 2005;89:724-733.
Not in our library10.1529/biophysj.105.060590
175.
Charras G.T., Yarrow J.C., Horton M.A., Mahadevan L., Mitchison T.J. Non-equilibration of hydrostatic pressure in blebbing cells. Nature. 2005;435:365-369.
Not in our library10.1038/nature03550
176.
Paluch E., Van Der Gucht J., Sykes C. Cracking up: Symmetry breaking in cellular systems. J. Cell Biol. 2006;175:687-692.
Not in our library10.1083/jcb.200607159
177.
Loitto V.M., Karlsson T., Magnusson K.E. Water flux in cell motility: Expanding the mechanisms of membrane protrusion. Cell Motil. Cytoskeleton. 2009;66:237-247.
Not in our library10.1002/cm.20357
178.
Blaser H., Reichman-Fried M., Castanon I., Dumstrei K., Marlow F.L., Kawakami K., Solnica-Krezel L., Heisenberg C.P., Raz E. Migration of Zebrafish Primordial Germ Cells: A Role for Myosin Contraction and Cytoplasmic Flow. Dev. Cell. 2006;11:613-627.
Not in our library10.1016/j.devcel.2006.09.023
179.
Sahai E., Marshall C.J. Differing modes for tumour cell invasion have distinct requirements for Rho/ROCK signalling and extracellular proteolysis. Nat. Cell Biol. 2003;5:711-719.
Not in our library10.1038/ncb1019
180.
Tournaviti S., Hannemann S., Terjung S., Kitzing T.M., Stegmayer C., Ritzerfeld J., Walther P., Grosse R., Nickel W., Fackler O.T. SH4-domain-induced plasma membrane dynamization promotes bleb-associated cell motility. J. Cell Sci. 2007;120:3820-3829.
Not in our library10.1242/jcs.011130
181.
Rossy J., Gutjahr M.C., Blaser N., Schlicht D., Niggli V. Ezrin/moesin in motile Walker 256 carcinosarcoma cells: Signal-dependent relocalization and role in migration. Exp. Cell Res. 2007;313:1106-1120.
Not in our library10.1016/j.yexcr.2006.12.023
182.
Brugues J., Maugis B., Casademunt J., Nassoy P., Amblard F., Sens P. Dynamical organization of the cytoskeletal cortex probed by micropipette aspiration. Proc. Natl. Acad. Sci. USA. 2010;107:15415-15420.
Not in our library10.1073/pnas.0913669107
183.
Charras G.T. A short history of blebbing. J. Microsc. 2008;231:466-478.
Not in our library10.1111/j.1365-2818.2008.02059.x
184.
Cunningham C.C. Actin polymerization and intracellular solvent flow in cell surfase blebbing. J. Cell Biol. 1995;129:1589-1599.
Not in our library10.1083/jcb.129.6.1589
185.
Strychalski W., Guy R.D. A computational model of bleb formation. Math. Med. Biol. 2013;30:115-130.
Not in our library10.1093/imammb/dqr030
186.
Keller H., Rentsch P., Hagmann J. Differences in cortical actin structure and dynamics document that different types of blebs are formed by distinct mechanisms. Exp. Cell Res. 2002;277:161-172.
Not in our library10.1006/excr.2002.5552
187.
Kristó I., Bajusz I., Bajusz C., Borkúti P., Vilmos P. Actin, actin-binding proteins, and actin-related proteins in the nucleus. Histochem. Cell Biol. 2016;145:373-388.
Not in our library10.1007/s00418-015-1400-9
188.
Bajusz C., Borkúti P., Kristó I., Kovács Z., Abonyi C., Vilmos P. Nuclear actin: Ancient clue to evolution in eukaryotes? Histochem. Cell Biol. 2018;150:1-10.
Not in our library10.1007/s00418-018-1693-6
189.
Le H.Q., Ghatak S., Yeung C.Y.C., Tellkamp F., Günschmann C., Dieterich C., Yeroslaviz A., Habermann B., Pombo A., Niessen C.M., et al. Mechanical regulation of transcription controls Polycomb-mediated gene silencing during lineage commitment. Nat. Cell Biol. 2016;18:864-875.
Not in our library10.1038/ncb3387
190.
Obrdlik A., Percipalle P. The F-actin severing protein cofilin-1 is required for RNA polymerase II transcription elongation. Nucleus. 2011;2:72-79.
Not in our library10.4161/nucl.14508
191.
Serebryannyy L.A., Parilla M., Annibale P., Cruz C.M., Laster K., Gratton E., Kudryashov D., Kosak S.T., Gottardi C.J., de Lanerolle P. Persistent nuclear actin filaments inhibit transcription by RNA polymerase II. J. Cell Sci. 2016;129:3412-3425.
Not in our library10.1242/jcs.195867
192.
Baarlink C., Plessner M., Sherrard A., Morita K., Misu S., Virant D., Kleinschnitz E.M., Harniman R., Alibhai D., Baumeister S., et al. A transient pool of nuclear F-actin at mitotic exit controls chromatin organization. Nat. Cell Biol. 2017;19:1389-1399.
Not in our library10.1038/ncb3641
193.
de Leeuw R., Gruenbaum Y., Medalia O. Nuclear Lamins: Thin Filaments with Major Functions. Trends Cell Biol. 2018;28:34-45.
Not in our library10.1016/j.tcb.2017.08.004
194.
Baarlink C., Wang H., Grosse R. Nuclear Actin Network Assembly by Formins Regulates the SRF Coactivator MAL. Science. 2013;2:864-868.
Not in our library10.1126/science.1235038
195.
Plessner M., Grosse R. Dynamizing nuclear actin filaments. Curr. Opin. Cell Biol. 2019;56:1-6.
Not in our library10.1016/j.ceb.2018.08.005
196.
Parisis N., Krasinska L., Harker B., Urbach S., Rossignol M., Camasses A., Dewar J., Morin N., Fisher D. Initiation of DNA replication requires actin dynamics and formin activity. EMBO J. 2017;e201796585.
Not in our library10.15252/embj.201796585
197.
DNA damage induces nuclear actin filament assembly by Formin-2 and Spire-1/2 that promotes efficient DNA repair · 2015
Corrected
198.
Caridi C.P., D’agostino C., Ryu T., Zapotoczny G., Delabaere L., Li X., Khodaverdian V.Y., Amaral N., Lin E., Rau A.R., et al. Nuclear F-actin and myosins drive relocalization of heterochromatic breaks. Nature. 2018;559:54-60.
Not in our library10.1038/s41586-018-0242-8
199.
Schrank B.R., Aparicio T., Li Y., Chang W., Chait B.T., Gundersen G.G., Gottesman M.E., Gautier J. Nuclear ARP2/3 drives DNA break clustering for homology-directed repair. Nature. 2018;559:61-66.
Not in our library10.1038/s41586-018-0237-5
200.
Kelpsch D.J., Tootle T.L. Nuclear Actin: From Discovery to Function. Anat. Rec. 2018;2013:1999-2013.
Not in our library10.1002/ar.23959
201.
Ohi R., Zanic M. Ahead of the Curve: New Insights into Microtubule Dynamics. F1000Research. 2016;5:314.
Not in our library10.12688/f1000research.7439.1
202.
Doxsey S. Re-evaluating centrosome function. Nat. Rev. Mol. Cell Biol. 2001;2:688-698.
Not in our library10.1038/35089575
203.
Vinogradova T., Miller P.M., Kaverina I. Microtubule network asymmetry in motile cells: Role of Golgi-derived array. Cell Cycle. 2009;8:2168-2174.
Not in our library10.4161/cc.8.14.9074
204.
Sanchez A.D., Feldman J.L. Microtubule-organizing centers: From the centrosome to non-centrosomal sites. Curr. Opin. Cell Biol. 2017;44:93-101.
Not in our library10.1016/j.ceb.2016.09.003
205.
Wu J., Akhmanova A. Microtubule-Organizing Centers. Annu. Rev. Cell Dev. Biol. 2017;33:51-75.
Not in our library10.1146/annurev-cellbio-100616-060615
206.
Dimitrov A., Quesnoit M., Moutel S., Cantaloube I., Poüs C., Perez F. Detection of GTP-tubulin conformation in vivo reveals a role for GTP remnants in microtubule rescues. Science. 2008;322:1353-1356.
Not in our library10.1126/science.1165401
207.
Howard J., Hyman A.A. Growth, fluctuation and switching at microtubule plus ends. Nat. Rev. Mol. Cell Biol. 2009;10:569-574.
Not in our library10.1038/nrm2713
208.
The size of the EB cap determines instantaneous microtubule stability · 2016
No notice
209.
Kita K., Wittmann T., Näthke I.S., Waterman-Storer C.M. Adenomatous polyposis coli on microtubule plus ends in cell extensions can promote microtubule net growth with or without EB1. Mol. Biol. Cell. 2006;17:2331-2345.
Not in our library10.1091/mbc.e05-06-0498
210.
Nakamura M., Zhou X.Z., Lu K.P. Critical role for the EB1 and APC interaction in the regulation of microtubule polymerization. Curr. Biol. 2001;11:1062-1067.
Not in our library10.1016/s0960-9822(01)00297-4
211.
Akhmanova A., Steinmetz M.O. Control of microtubule organization and dynamics: Two ends in the limelight. Nat. Rev. Mol. Cell Biol. 2015;16:711-726.
Not in our library10.1038/nrm4084
212.
Mimori-Kiyosue Y., Grigoriev I., Lansbergen G., Sasaki H., Matsui C., Severin F., Galjart N., Grosveld F., Vorobjev I., Tsukita S., et al. CLASP1 and CLASP2 bind to EB1 and regulate microtubule plus-end dynamics at the cell cortex. J. Cell Biol. 2005;168:141-153.
Not in our library10.1083/jcb.200405094
213.
Suozzi K.C., Wu X., Fuchs E. Spectraplakins: Master orchestrators of cytoskeletal dynamics. J. Cell Biol. 2012;197:465-475.
Not in our library10.1083/jcb.201112034
214.
Drabek K., van Ham M., Stepanova T., Draegestein K., van Horssen R., Sayas C.L., Akhmanova A., ten Hagen T., Smits R., Fodde R., et al. Role of CLASP2 in Microtubule Stabilization and the Regulation of Persistent Motility. Curr. Biol. 2006;16:2259-2264.
Not in our library10.1016/j.cub.2006.09.065
215.
Leung C.L., Sun D., Zheng M., Knowles D.R., Liem R.K. Microtubule actin cross-linking factor (MACF): A hybrid of dystonin and dystrophin that can interact with the actin and microtubule cytoskeletons. J. Cell Biol. 1999;147:1275-1286.
Not in our library10.1083/jcb.147.6.1275
216.
Komarova Y., De Groot C.O., Grigoriev I., Gouveia S.M., Munteanu E.L., Schober J.M., Honnappa S., Buey R.M., Hoogenraad C.C., Dogterom M., et al. Mammalian end binding proteins control persistent microtubule growth. J. Cell Biol. 2009;184:691-706.
Not in our library10.1083/jcb.200807179
217.
Maurer S.P., Cade N.I., Bohner G., Gustafsson N., Boutant E., Surrey T. EB1 Accelerates Two Conformational Transitions Important for Microtubule Maturation and Dynamics. Curr. Biol. 2014;24:372-384.
Not in our library10.1016/j.cub.2013.12.042
218.
Dong Z., Zhou L., Han N., Zhang M., Lyu X. Wnt/β-catenin pathway involvement in ionizing radiation-induced invasion of U87 glioblastoma cells. Strahlentherapie und Onkol. 2015;191:672-680.
Not in our library10.1007/s00066-015-0858-7
219.
Evans K.J., Gomes E.R., Reisenweber S.M., Gundersen G.G., Lauring B.P. Linking axonal degeneration to microtubule remodeling by Spastin-mediated microtubule severing. J. Cell Biol. 2005;168:599-606.
Not in our library10.1083/jcb.200409058
220.
Sharp D.J., Ross J.L. Microtubule-severing enzymes at the cutting edge. J. Cell Sci. 2012;125:2561-2569.
Not in our library10.1242/jcs.101139
221.
Gupta K.K., Li C., Duan A., Alberico E.O., Kim O.V., Alber M.S., Goodson H. V Mechanism for the catastrophe-promoting activity of the microtubule destabilizer Op18/stathmin. Proc. Natl. Acad. Sci. USA. 2013;110:20449-20454.
Not in our library10.1073/pnas.1309958110
222.
Brouhard G.J., Stear J.H., Noetzel T.L., Al-Bassam J., Kinoshita K., Harrison S.C., Howard J., Hyman A.A. XMAP215 Is a Processive Microtubule Polymerase. Cell. 2008;132:79-88.
Not in our library10.1016/j.cell.2007.11.043
223.
Moores C.A., Perderiset M., Kappeler C., Kain S., Drummond D., Perkins S.J., Chelly J., Cross R., Houdusse A., Francis F. Distinct roles of doublecortin modulating the microtubule cytoskeleton. EMBO J. 2006;25:4448-4457.
Not in our library10.1038/sj.emboj.7601335
224.
Dehmelt L., Halpain S. The MAP2/Tau family of microtubule-associated proteins. Genome Biol. 2004;6:204.
Not in our library10.1186/gb-2004-6-1-204
225.
Qiang L., Yu W., Andreadis A., Luo M., Baas P.W. Tau Protects Microtubules in the Axon from Severing by Katanin. J. Neurosci. 2006;26:3120-3129.
Not in our library10.1523/jneurosci.5392-05.2006
226.
Miller P.M., Folkmann A.W., Maia A.R.R., Efimova N., Efimov A., Kaverina I. Golgi-derived CLASP-dependent microtubules control Golgi organization and polarized trafficking in motile cells. Nat. Cell Biol. 2009;11:1069-1080.
Not in our library10.1038/ncb1920
227.
Yadav S., Puri S., Linstedt A. A Primary Role for Golgi Positioning in Directed Secretion, Cell Polarity, and Wound Healing. Mol. Biol. Cell. 2009;20:1728-1736.
Not in our library10.1091/mbc.e08-10-1077
228.
Verhey K.J., Kaul N., Soppina V. Kinesin Assembly and Movement in Cells. Annu. Rev. Biophys. 2011;40:267-288.
Not in our library10.1146/annurev-biophys-042910-155310
229.
Cianfrocco M.A., DeSantis M.E., Leschziner A.E., Reck-Peterson S.L. Mechanism and regulation of cytoplasmic dynein. Annu. Rev. Cell Dev. Biol. 2015;31:83-108.
Not in our library10.1146/annurev-cellbio-100814-125438
230.
Osmani N., Peglion F., Chavrier P., Etienne-Manneville S. Cdc42 localization and cell polarity depend on membrane traffic. J. Cell Biol. 2010;191:1261-1269.
Not in our library10.1083/jcb.201003091
231.
Palamidessi A., Frittoli E., Garré M., Faretta M., Mione M., Testa I., Diaspro A., Lanzetti L., Scita G., Di Fiore P.P. Endocytic Trafficking of Rac Is Required for the Spatial Restriction of Signaling in Cell Migration. Cell. 2008;134:135-147.
Not in our library10.1016/j.cell.2008.05.034
232.
Hookway C., Ding L., Davidson M.W., Rappoport J.Z., Danuser G., Gelfand V.I. Microtubule-dependent transport and dynamics of vimentin intermediate filaments. Mol. Biol. Cell. 2015;26:1675-1686.
Not in our library10.1091/mbc.e14-09-1398
233.
Robert A., Tian P., Adam S.A., Kittisopikul M., Jaqaman K., Goldman R.D., Gelfand V.I. Kinesin-dependent transport of keratin filaments: A unified mechanism for intermediate filament transport. FASEB J. 2018;fj.201800604R.
Not in our library10.1096/fj.201800604r
234.
Helfand B.T., Mikami A., Vallee R.B., Goldman R.D. A requirement for cytoplasmic dynein and dynactin in intermediate filament network assembly and organization. J. Cell Biol. 2002;157:795-806.
Not in our library10.1083/jcb.200202027
235.
Mingle L.A. Localization of all seven messenger RNAs for the actin-polymerization nucleator Arp2/3 complex in the protrusions of fibroblasts. J. Cell Sci. 2005;118:2425-2433.
Not in our library10.1242/jcs.02371
236.
Oleynikov Y., Singer R.H. RNA localization: Different zipcodes, same postman? Trends Cell Biol. 1998;8:381-383.
Not in our library10.1016/s0962-8924(98)01348-8
237.
Su X., Ohi R., Pellman D. Move in for the kill: Motile microtubule regulators. Trends Cell Biol. 2012;22:567-575.
Not in our library10.1016/j.tcb.2012.08.003
238.
Moores C.A., Yu M., Guo J., Beraud C., Sakowicz R., Milligan R.A. A Mechanism for Microtubule Depolymerization by KinI Kinesins. Mol. Cell. 2002;9:903-909.
Not in our library10.1016/s1097-2765(02)00503-8
239.
Liu M., Nadar V.C., Kozielski F., Kozlowska M., Yu W., Baas P.W. Kinesin-12, a mitotic microtubule-associated motor protein, impacts axonal growth, navigation, and branching. J. Neurosci. 2010;30:14896-14906.
Not in our library10.1523/jneurosci.3739-10.2010
240.
Myers K.A., Baas P.W. Kinesin-5 regulates the growth of the axon by acting as a brake on its microtubule array. J. Cell Biol. 2007;178:1081-1091.
Not in our library10.1083/jcb.200702074
241.
Jolly A.L., Kim H., Srinivasan D., Lakonishok M., Larson A.G., Gelfand V.I. Kinesin-1 heavy chain mediates microtubule sliding to drive changes in cell shape. Proc. Natl. Acad. Sci. USA. 2010;107:12151-12156.
Not in our library10.1073/pnas.1004736107
242.
Song Y., Brady S.T. Posttranslational Modifications of Tubulin: Pathways to Functional Diversity of Microtubules. Trends Cell Biol. 2015;25:125-136.
Not in our library10.1016/j.tcb.2014.10.004
243.
Portran D., Schaedel L., Xu Z., Théry M., Nachury M.V. Tubulin acetylation protects long-lived microtubules against mechanical ageing. Nat. Cell Biol. 2017;19:391-398.
Not in our library10.1038/ncb3481
244.
Sudo H., Baas P.W. Acetylation of microtubules influences their sensitivity to severing by katanin in neurons and fibroblasts. J. Neurosci. 2010;30:7215-7226.
Not in our library10.1523/jneurosci.0048-10.2010
245.
Roll-Mecak A. Intrinsically disordered tubulin tails: Complex tuners of microtubule functions? Semin. Cell Dev. Biol. 2015;37:11-19.
Not in our library10.1016/j.semcdb.2014.09.026
246.
Konishi Y., Setou M. Tubulin tyrosination navigates the kinesin-1 motor domain to axons. Nat. Neurosci. 2009;12:559-567.
Not in our library10.1038/nn.2314
247.
Roll-Mecak A., Vale R.D. Structural basis of microtubule severing by the hereditary spastic paraplegia protein spastin. Nature. 2008;451:363-367.
Not in our library10.1038/nature06482
248.
Lacroix B., Van Dijk J., Gold N.D., Guizetti J., Aldrian-Herrada G., Rogowski K., Gerlich D.W., Janke C. Tubulin polyglutamylation stimulates spastin-mediated microtubule severing. J. Cell Biol. 2010;189:945-954.
Not in our library10.1083/jcb.201001024
249.
Valenstein M.L., Roll-Mecak A. Graded Control of Microtubule Severing by Tubulin Glutamylation. Cell. 2016;164:911-921.
Not in our library10.1016/j.cell.2016.01.019
250.
Morris E.J., Nader G.P.F., Ramalingam N., Bartolini F., Gundersen G.G. Kif4 interacts with EB1 and stabilizes microtubules downstream of Rho-mDia in migrating fibroblasts. PLoS ONE. 2014;9.
Not in our library10.1371/journal.pone.0091568
251.
Wen Y., Eng C.H., Schmoranzer J., Cabrera-Poch N., Morris E.J.S., Chen M., Wallar B.J., Alberts A.S., Gundersen G.G. EB1 and APC bind to mDia to EB1 and APC bind to mDia to stabilize stabilize microtubules downstream of Rho and promote cell migration. Nat. Cell Biol. 2004;6:820-830.
Not in our library10.1038/ncb1160
252.
Ishizaki T., Morishima Y., Okamoto M., Furuyashiki T., Kato T., Narumiya S. Coordination of microtubules and the actin cytoskeleton by the Rho effector mDia1. Nat. Cell Biol. 2001;3:8-14.
Not in our library10.1038/35050598
253.
Fife C.M., Sagnella S.M., Teo W.S., Po’Uha S.T., Byrne F.L., Yeap Y.Y.C., Ng D.C.H., Davis T.P., McCarroll J.A., Kavallaris M. Stathmin mediates neuroblastoma metastasis in a tubulin-independent manner via RhoA/ROCK signaling and enhanced transendothelial migration. Oncogene. 2017;36:501-511.
Not in our library10.1038/onc.2016.220
254.
Daub H., Gevaert K., Vandekerckhove J., Sobel A., Hall A. Rac/Cdc42 and p65PAK regulate the microtubule-destabilizing protein stathmin through phosphorylation at serine 16. J. Biol. Chem. 2001;276:1677-1680.
Not in our library10.1074/jbc.c000635200
255.
Wittmann T., Bokoch G.M., Waterman-Storer C.M. Regulation of microtubule destabilizing activity of Op18/stathmin downstream of Rac1. J. Biol. Chem. 2004;279:6196-6203.
Not in our library10.1074/jbc.m307261200
256.
Wittmann T., Bokoch G.M., Waterman-Storer C.M. Regulation of leading edge microtubule and actin dynamics downstream of Rac1. J. Cell Biol. 2003;161:845-851.
Not in our library10.1083/jcb.200303082
257.
Fukata M., Watanabe T., Noritake J., Nakagawa M., Yamaga M., Kuroda S., Matsuura Y., Iwamatsu A., Perez F., Kaibuchi K. Rac1 and Cdc42 capture microtubules through IQGAP1 and CLIP-170. Cell. 2002;109:873-885.
Not in our library10.1016/s0092-8674(02)00800-0
258.
Brown R.A., Talas G., Porter R.A., McGrouther D.A., Eastwood M. Balanced mechanical forces and microtubule contribution to fibroblast contraction. J. Cell. Physiol. 1996;169:439-447.
Not in our library10.1002/(sici)1097-4652(199612)169:3<439::aid-jcp4>3.0.co;2-p
259.
Rudolph R., Woodward M. Spatial orientation of microtubules in contractile fibroblasts in vivo. Anat. Rec. 1978;191:169-181.
Not in our library10.1002/ar.1091910204
260.
Tomasek J.J., Hay E.D. Analysis of the role of microfilaments and microtubules in acquisition of bipolarity and elongation of fibroblasts in hydrated collagen gels. J. Cell Biol. 1984;99:536-549.
Not in our library10.1083/jcb.99.2.536
261.
Etienne-Manneville S. Microtubules in Cell Migration. Annu. Rev. Cell Dev. Biol. 2013;29:471-499.
Not in our library10.1146/annurev-cellbio-101011-155711
262.
Hawkins T., Mirigian M., Selcuk Yasar M., Ross J.L. Mechanics of microtubules. J. Biomech. 2010;43:23-30.
Not in our library10.1016/j.jbiomech.2009.09.005
263.
Dogterom M., Yurke B. Measurement of the force-velocity relation for growing microtubules. Science. 1997;278:856-860.
Not in our library10.1126/science.278.5339.856
264.
Kolomeisky A.B., Fisher M.E. Force-velocity relation for growing microtubules. Biophys. J. 2001;80:149-154.
Not in our library10.1016/s0006-3495(01)76002-x
265.
Janson M.E., de Dood M.E., Dogterom M. Dynamic instability of microtubules is regulated by force. J. Cell Biol. 2003;161:1029-1034.
Not in our library10.1083/jcb.200301147
266.
Robison P., Caporizzo M.A., Ahmadzadeh H., Bogush A.I., Chen C.Y., Margulies K.B., Shenoy V.B., Prosser B.L. Detyrosinated microtubules buckle and bear load in contracting cardiomyocytes. Science. 2016;352:aaf0659.
Not in our library10.1126/science.aaf0659
267.
Brangwynne C.P., MacKintosh F.C., Kumar S., Geisse N.A., Talbot J., Mahadevan L., Parker K.K., Ingber D.E., Weitz D.A. Microtubules can bear enhanced compressive loads in living cells because of lateral reinforcement. J. Cell Biol. 2006;173:733-741.
Not in our library10.1083/jcb.200601060
268.
Laan L., Husson J., Munteanu E.L., Kerssemakers J.W.J., Dogterom M. Force-generation and dynamic instability of microtubule bundles. Proc. Natl. Acad. Sci. USA. 2008;105:8920-8925.
Not in our library10.1073/pnas.0710311105
269.
Komarova Y.A., Vorobjev I.A., Borisy G.G. Life cycle of MTs: Persistent growth in the cell interior, asymmetric transition frequencies and effects of the cell boundary. J. Cell Sci. 2002;115:3527-3539.
Not in our library10.1242/jcs.115.17.3527
270.
Drummond D.R., Cross R.A. Dynamics of interphase microtubules in Schizosaccharomyces pombe. Curr. Biol. 2000;10:766-775.
Not in our library10.1016/s0960-9822(00)00570-4
271.
Bicek A.D., Tüzel E., Demtchouk A., Uppalapati M., Hancock W.O., Kroll D.M., Odde D.J. Anterograde microtubule transport drives microtubule bending in LLC-PK1 epithelial cells. Mol. Biol. Cell. 2009;20:2943-2953.
Not in our library10.1091/mbc.e08-09-0909
272.
Desai A., Mitchison T.J. Microtubule Polymerization Dynamics. Annu. Rev. Cell Dev. Biol. 1997;13:83-117.
Not in our library10.1146/annurev.cellbio.13.1.83
273.
Grishchuk E.L., Molodtsov M.I., Ataullakhanov F.I., McIntosh J.R. Force production by disassembling microtubules. Nature. 2005;438:384-388.
Not in our library10.1038/nature04132
274.
Petry S. Mechanisms of Mitotic Spindle Assembly. Annu. Rev. Biochem. 2016;85:659-683.
Not in our library10.1146/annurev-biochem-060815-014528
275.
Duro E., Marston A.L. From equator to pole: Splitting chromosomes in mitosis and meiosis. Genes Dev. 2015;29:109-122.
Not in our library10.1101/gad.255554.114
276.
Gadde S., Heald R. Mechanisms and Molecules of the Mitotic Spindle. Curr. Biol. 2004;14:R797-R805.
Not in our library10.1016/j.cub.2004.09.021
277.
Okada K., Bartolini F., Deaconescu A.M., Moseley J.B., Dogic Z., Grigorieff N., Gundersen G.G., Goode B.L. Adenomatous polyposis coli protein nucleates actin assembly and synergizes with the formin mDia1. J. Cell Biol. 2010;189:1087-1096.
Not in our library10.1083/jcb.201001016
278.
Bartolini F., Moseley J.B., Schmoranzer J., Cassimeris L., Goode B.L., Gundersen G.G. The formin mDia2 stabilizes microtubules independently of its actin nucleation activity. J. Cell Biol. 2008;181:523-536.
Not in our library10.1083/jcb.200709029
279.
Wu X., Kodama A., Fuchs E. ACF7 Regulates Cytoskeletal-Focal Adhesion Dynamics and Migration and Has ATPase Activity. Cell. 2008;135:137-148.
Not in our library10.1016/j.cell.2008.07.045
280.
Miller A.L., Wang Y., Mooseker M.S., Koleske A.J. The Abl-related gene (Arg) requires its F-actin–microtubule cross-linking activity to regulate lamellipodial dynamics during fibroblast adhesion. J. Cell Biol. 2004;165:407-420.
Not in our library10.1083/jcb.200308055
281.
Moll R., Divo M., Langbein L. The human keratins: Biology and pathology. Histochem. Cell Biol. 2008;129:705-733.
Not in our library10.1007/s00418-008-0435-6
282.
Eliasson C., Sahlgren C., Berthold C.H., Stakeberg J., Celis J.E., Betsholtz C., Eriksson J.E., Pekny M. Intermediate filament protein partnership in astrocytes. J. Biol. Chem. 1999;274:23996-24006.
Not in our library10.1074/jbc.274.34.23996
283.
Bellin R.M., Sernett S.W., Becker B., Ip W., Huiatt T.W., Robson R.M. Molecular characteristics and interactions of the intermediate filament protein synemin. Interactions with alpha-actinin may anchor synemin-containing heterofilaments. J. Biol. Chem. 1999;274:29493-29499.
Not in our library10.1074/jbc.274.41.29493
284.
Herrmann H., Aebi U. Intermediate Filaments: Molecular Structure, Assembly Mechanism, and Integration Into Functionally Distinct Intracellular Scaffolds. Annu. Rev. Biochem. 2004;73:749-789.
Not in our library10.1146/annurev.biochem.73.011303.073823
285.
Fuchs E. Intermediate Filaments: Structure, Dynamics, Function, and Disease. Annu. Rev. Biochem. 1994;63:345-382.
Not in our library10.1146/annurev.bi.63.070194.002021
286.
Smith T.A., Strelkov S.V., Burkhard P., Aebi U., Parry D.A.D. Sequence comparisons of intermediate filament chains: Evidence of a unique functional/structural role for coiled-coil segment 1A and linker L1. J. Struct. Biol. 2002;137:128-145.
Not in our library10.1006/jsbi.2002.4438
287.
Sokolova A.V., Kreplak L., Wedig T., Mucke N., Svergun D.I., Herrmann H., Aebi U., Strelkov S.V. Monitoring intermediate filament assembly by small-angle x-ray scattering reveals the molecular architecture of assembly intermediates. Proc. Natl. Acad. Sci. USA. 2006;103:16206-16211.
Not in our library10.1073/pnas.0603629103
288.
Herrmann H., Häner M., Brettel M., Ku N.O., Aebi U. Characterization of distinct early assembly units of different intermediate filament proteins. J. Mol. Biol. 1999;286:1403-1420.
Not in our library10.1006/jmbi.1999.2528
289.
Mücke N., Wedig T., Bürer A., Marekov L.N., Steinert P.M., Langowski J., Aebi U., Herrmann H. Molecular and biophysical characterization of assembly-starter units of human vimentin. J. Mol. Biol. 2004;340:97-114.
Not in our library10.1016/j.jmb.2004.04.039
290.
Georgakopoulou S., Möller D., Sachs N., Herrmann H., Aebi U. Near-UV Circular Dichroism Reveals Structural Transitions of Vimentin Subunits during Intermediate Filament Assembly. J. Mol. Biol. 2009;386:544-553.
Not in our library10.1016/j.jmb.2008.12.053
291.
Steinert P.M., Marekov L.N., Parry D.A. Diversity of intermediate filament structure. J. Biol. Chem. 1993;268:24916-24925.
Not in our library
292.
Herrmann H., Kreplak L., Aebi U. Isolation, Characterization, and In Vitro Assembly of Intermediate Filaments. Methods Cell Biol. 2004;78:3-24.
Not in our library10.1016/s0091-679x(04)78001-2
293.
Nöding B., Herrmann H., Köster S. Direct Observation of Subunit Exchange along Mature Vimentin Intermediate Filaments. Biophys. J. 2014;107:2923-2931.
Not in our library10.1016/j.bpj.2014.09.050
294.
Chang L., Goldman R.D. Intermediate filaments mediate cytoskeletal crosstalk. Mol. Cell Biol. 2004;5:601-613.
Not in our library10.1038/nrm1438
295.
Kayser J., Haslbeck M., Dempfle L., Krause M., Grashoff C., Buchner J., Herrmann H., Bausch A.R. The Small Heat Shock Protein Hsp27 Affects Assembly Dynamics and Structure of Keratin Intermediate Filament Networks. Biophys. J. 2013;105:1778-1785.
Not in our library10.1016/j.bpj.2013.09.007
296.
Heitlinger E., Peter M., Lustig A., Villiger W., Nigg E.A., Aebi U. The role of the head and tail domain in lamin structure and assembly: Analysis of bacterially expressed chicken Lamin A and truncated B2 lamins. J. Struct. Biol. 1992;108:74-91.
Not in our library10.1016/1047-8477(92)90009-y
297.
Heitlinger E., Peter M., Häner M., Lustig A., Aebi U., Nigg E.A. Expression of chicken lamin B2 in Escherichia coli: Characterization of its structure, assembly, and molecular interactions. J. Cell Biol. 1991;113:485-495.
Not in our library10.1083/jcb.113.3.485
298.
Aebi U., Cohn J., Buhle L., Gerace L. The nuclear lamina is a meshwork of intermediate-type filaments. Nature. 1986;323:560-564.
Not in our library10.1038/323560a0
299.
Kreis S., Schönfeld H.J., Melchior C., Steiner B., Kieffer N. The intermediate filament protein vimentin binds specifically to a recombinant integrin α2/β1 cytoplasmic tail complex and co-localizes with native α2/β1 in endothelial cell focal adhesions. Exp. Cell Res. 2005;305:110-121.
Not in our library10.1016/j.yexcr.2004.12.023
300.
Tsuruta D. The vimentin cytoskeleton regulates focal contact size and adhesion of endothelial cells subjected to shear stress. J. Cell Sci. 2003;116:4977-4984.
Not in our library10.1242/jcs.00823
301.
Seltmann K., Cheng F., Wiche G., Eriksson J.E., Magin T.M. Keratins stabilize hemidesmosomes through regulation of β4-integrin turnover. J. Invest. Dermatol. 2015;135:1609-1620.
Not in our library10.1038/jid.2015.46
302.
Loschke F., Homberg M., Magin T.M. Keratin isotypes control desmosome stability and dynamics through PKCα. J. Invest. Dermatol. 2016;136:202-213.
Not in our library10.1038/jid.2015.403
303.
Etienne-Manneville S. Cytoplasmic Intermediate Filaments in Cell Biology. Annu. Rev. Cell Dev. Biol. 2018;34:1-28.
Not in our library10.1146/annurev-cellbio-100617-062534
304.
Dupin I., Sakamoto Y., Etienne-Manneville S. Cytoplasmic intermediate filaments mediate actin-driven positioning of the nucleus. J. Cell Sci. 2011;124:865-872.
Not in our library10.1242/jcs.076356
305.
Matveeva E.A., Venkova L.S., Chernoivanenko I.S., Minin A.A. Vimentin is involved in regulation of mitochondrial motility and membrane potential by Rac1. Biol. Open. 2015;4:1290-1297.
Not in our library10.1242/bio.011874
306.
Nekrasova O.E., Mendez M.G., Chernoivanenko I.S., Tyurin-Kuzmin P.A., Kuczmarski E.R., Gelfand V.I., Goldman R.D., Minin A.A. Vimentin intermediate filaments modulate the motility of mitochondria. Mol. Biol. Cell. 2011;22:2282-2289.
Not in our library10.1091/mbc.e10-09-0766
307.
Wang N., Stamenović D. Contribution of intermediate filaments to cell stiffness, stiffening, and growth. Am. J. Physiol. Cell Physiol. 2000;279:C188-C194.
Not in our library10.1152/ajpcell.2000.279.1.c188
308.
Godsel L.M., Hobbs R.P., Green K.J. Intermediate filament assembly: Dynamics to disease. Trends Cell Biol. 2008;18:28-37.
Not in our library10.1016/j.tcb.2007.11.004
309.
Intermediate Filaments at the Junction of Mechanotransduction, Migration, and Development · 2017
No notice
310.
Storm C., Pastore J.J., Mackintosh F.C., Lubensky T.C., Janmey P.A. Nonlinear elasticity in biological gels. Nature. 2005;435:191-194.
Not in our library10.1038/nature03521
311.
Guo M., Ehrlicher A.J., Mahammad S., Fabich H., Jensen M.H., Moore J.R., Fredberg J.J., Goldman R.D., Weitz D.A. The Role of Vimentin Intermediate Filaments in Cortical and Cytoplasmic Mechanics. Biophys. J. 2013;105:1562-1568.
Not in our library10.1016/j.bpj.2013.08.037
312.
Mendez M.G., Restle D., Janmey P.A. Vimentin enhances cell elastic behavior and protects against compressive stress. Biophys. J. 2014;107:314-323.
Not in our library10.1016/j.bpj.2014.04.050
313.
Vimentin contributes to epithelial-mesenchymal transition cancer cell mechanics by mediating cytoskeletal organization and focal adhesion maturation · 2015
No notice
314.
Alberti C. Cytoskeleton structure and dynamic behaviour: Quick excursus from basic molecular mechanisms to some implications in cancer chemotherapy. Eur. Rev. Med. Pharmacol. Sci. 2009;13:13-21.
Not in our library
315.
Izawa I., Inagaki M. Regulatory mechanisms and functions of intermediate filaments: A study using site- and phosphorylation state-specific antibodies. Cancer Sci. 2006;97:167-174.
Not in our library10.1111/j.1349-7006.2006.00161.x
316.
Hyder C.L., Pallari H.-M., Kochin V., Eriksson J.E. Providing cellular signposts - Post-translational modifications of intermediate filaments. FEBS Lett. 2008;582:2140-2148.
Not in our library10.1016/j.febslet.2008.04.064
317.
Pan X., Hobbs R.P., Coulombe P.A. The expanding significance of keratin intermediate filaments in normal and diseased epithelia. Curr. Opin. Cell Biol. 2013;25:47-56.
Not in our library10.1016/j.ceb.2012.10.018
318.
Rogel M.R., Jaitovich A., Ridge K.M. The Role of the Ubiquitin Proteasome Pathway in Keratin Intermediate Filament Protein Degradation. Proc. Am. Thorac. Soc. 2010;7:71-76.
Not in our library10.1513/pats.200908-089js
319.
Omary M.B., Coulombe P.A., McLean W.H.I. Intermediate Filament Proteins and Their Associated Diseases. N. Engl. J. Med. 2004;351:2087-2100.
Not in our library10.1056/nejmra040319
320.
Leung C.L., Liem R.K.H., Parry D.A.D., Green K.J. The plakin family. J. Cell Sci. 2001;114:3409-3410.
Not in our library10.1242/jcs.114.19.3409
321.
Leung C.L., Green K.J., Liem R.K.H. Plakins: A family of versatile cytolinker proteins. Trends Cell Biol. 2002;12:37-45.
Not in our library10.1016/s0962-8924(01)02180-8
322.
Desai B.V., Harmon R.M., Green K.J. Desmosomes at a glance. J. Cell Sci. 2009;122:4401-4407.
Not in our library10.1242/jcs.037457
323.
Wiche G., Osmanagic-Myers S., Castañón M.J. Networking and anchoring through plectin: A key to IF functionality and mechanotransduction. Curr. Opin. Cell Biol. 2015;32:21-29.
Not in our library10.1016/j.ceb.2014.10.002
324.
Kölsch A., Windoffer R., Leube R.E. Actin-dependent dynamics of keratin filament precursors. Cell Motil. Cytoskeleton. 2009;66:976-985.
Not in our library10.1002/cm.20395
325.
Robert A., Herrmann H., Davidson M.W., Gelfand V.I. Microtubule-dependent transport of vimentin filament precursors is regulated by actin and by the concerted action of Rho- and p21-activated kinases. FASEB J. 2014;28:2879-2890.
Not in our library10.1096/fj.14-250019
326.
Burgstaller G., Gregor M., Winter L., Wiche G. Keeping the vimentin network under control: Cell-matrix adhesion-associated plectin 1f affects cell shape and polarity of fibroblasts. Mol. Biol. Cell. 2010;21:3362-3375.
Not in our library10.1091/mbc.e10-02-0094
327.
Lynch C.D., Lazar A.M., Iskratsch T., Zhang X., Sheetz M.P. Endoplasmic spreading requires coalescence of vimentin intermediate filaments at force-bearing adhesions. Mol. Biol. Cell. 2013;24:21-30.
Not in our library10.1091/mbc.e12-05-0377
328.
Gregor M., Osmanagic-Myers S., Burgstaller G., Wolfram M., Fischer I., Walko G., Resch G.P., Jörgl A., Herrmann H., Wiche G. Mechanosensing through focal adhesion-anchored intermediate filaments. FASEB J. 2014;28:715-729.
Not in our library10.1096/fj.13-231829
329.
Esue O., Carson A.A., Tseng Y., Wirtz D. A direct interaction between actin and vimentin filaments mediated by the tail domain of vimentin. J. Biol. Chem. 2006;281:30393-30399.
Not in our library10.1074/jbc.m605452200
330.
Mellad J.A., Warren D.T., Shanahan C.M. Nesprins LINC the nucleus and cytoskeleton. Curr. Opin. Cell Biol. 2011;23:47-54.
Not in our library10.1016/j.ceb.2010.11.006
331.
Wilhelmsen K., Litjens S.H.M., Kuikman I., Tshimbalanga N., Janssen H., van den Bout I., Raymond K., Sonnenberg A. Nesprin-3, a novel outer nuclear membrane protein, associates with the cytoskeletal linker protein plectin. J. Cell Biol. 2005;171:799-810.
Not in our library10.1083/jcb.200506083
332.
Lombardi M.L., Jaalouk D.E., Shanahan C.M., Burke B., Roux K.J., Lammerding J. The Interaction between Nesprins and Sun Proteins at the Nuclear Envelope Is Critical for Force Transmission between the Nucleus and Cytoskeleton. J. Biol. Chem. 2011;286:26743-26753.
Not in our library10.1074/jbc.m111.233700
333.
Morgan J.T., Pfeiffer E.R., Thirkill T.L., Kumar P., Peng G., Fridolfsson H.N., Douglas G.C., Starr D.A., Barakat A.I. Nesprin-3 regulates endothelial cell morphology, perinuclear cytoskeletal architecture, and flow-induced polarization. Mol. Biol. Cell. 2011;22:4324-4334.
Not in our library10.1091/mbc.e11-04-0287
334.
Keeling M.C., Flores L.R., Dodhy A.H., Murray E.R., Gavara N. Actomyosin and vimentin cytoskeletal networks regulate nuclear shape, mechanics and chromatin organization. Sci. Rep. 2017;7:1-14.
Not in our library10.1038/s41598-017-05467-x
335.
Toivola D.M., Tao G.Z., Habtezion A., Liao J., Omary M.B. Cellular integrity plus: Organelle-related and protein-targeting functions of intermediate filaments. Trends Cell Biol. 2005;15:608-617.
Not in our library10.1016/j.tcb.2005.09.004
336.
Naetar N., Ferraioli S., Foisner R. Lamins in the nuclear interior—Life outside the lamina. J. Cell Sci. 2017;130:2087-2096.
Not in our library10.1242/jcs.203430
337.
Turgay Y., Eibauer M., Goldman A.E., Shimi T., Khayat M., Ben-Harush K., Dubrovsky-Gaupp A., Sapra K.T., Goldman R.D., Medalia O. The molecular architecture of lamins in somatic cells. Nature. 2017;543:261-264.
Not in our library10.1038/nature21382
338.
Buxboim A., Swift J., Irianto J., Spinler K.R., Dingal P.C.D.P., Athirasala A., Kao Y.C., Cho S., Harada T., Shin J.-W., et al. Matrix elasticity regulates lamin-A,C phosphorylation and turnover with feedback to actomyosin. Curr. Biol. 2015;24:1909-1917.
Not in our library10.1016/j.cub.2014.07.001
339.
Davidson P.M., Denais C., Bakshi M.C., Lammerding J. Nuclear deformability constitutes a rate-limiting step during cell migration in 3-D environments. Cell. Mol. Bioeng. 2014;7:293-306.
Not in our library10.1007/s12195-014-0342-y
340.
Harada T., Swift J., Irianto J., Shin J.W., Spinler K.R., Athirasala A., Diegmiller R., Dingal P.C.D.P., Ivanovska I.L., Discher D.E. Nuclear lamin stiffness is a barrier to 3D migration, but softness can limit survival. J. Cell Biol. 2014;204:669-682.
Not in our library10.1083/jcb.201308029
341.
Cho S., Irianto J., Discher D.E. Mechanosensing by the nucleus: From pathways to scaling relationships. J. Cell Biol. 2017;1-11.
Not in our library10.1083/jcb.201610042
342.
Osmanagic-Myers S., Dechat T., Foisner R. Lamins at the crossroads of mechanosignaling. Genes Dev. 2015;29:225-237.
Not in our library10.1101/gad.255968.114
343.
Stephens A.D., Banigan E.J., Adam S.A., Goldman R.D., Marko J.F. Chromatin and lamin A determine two different mechanical response regimes of the cell nucleus. Mol. Biol. Cell. 2017;28:1984-1996.
Not in our library10.1091/mbc.e16-09-0653
344.
Vicente-Manzanares M., Webb D.J., Horwitz A.R. Cell migration at a glance. J. Cell Sci. 2005;118:4917-4919.
Not in our library10.1242/jcs.02662
345.
Friedl P., Wolf K. Plasticity of cell migration: A multiscale tuning model. J. Cell Biol. 2010;188:11-19.
Not in our library10.1083/jcb.200909003
346.
Rac and Rho GTPases in cancer cell motility control · 2010
No notice
347.
Nguyen L.K., Kholodenko B.N., von Kriegsheim A. Rac1 and RhoA: Networks, loops and bistability. Small GTPases. 2016;0:1-6.
Not in our library10.1080/21541248.2016.1224399
348.
Brunner C.A., Ehrlicher A., Kohlstrunk B., Knebel D., Käs J.A., Goegler M. Cell migration through small gaps. Eur. Biophys. J. 2006;35:713-719.
Not in our library10.1007/s00249-006-0079-1
349.
Heinemann F., Doschke H., Radmacher M. Keratocyte lamellipodial protrusion is characterized by a concave force-velocity relation. Biophys. J. 2011;100:1420-1427.
Not in our library10.1016/j.bpj.2011.01.063
350.
Pollard T.D., Borisy G.G. Cellular motility driven by assembly and disassembly of actin filaments. Cell. 2003;112:453-465.
Not in our library10.1016/s0092-8674(03)00120-x
351.
FMNL formins boost lamellipodial force generation · 2017
No notice
352.
Gorelik R., Gautreau A. The Arp2/3 inhibitory protein arpin induces cell turning by pausing cell migration. Cytoskeleton. 2015;72:362-371.
Not in our library10.1002/cm.21233
353.
Choi C.K., Vicente-Manzanaras M., Zareno J., Whitmore L.A., Mogilner A., Horwitz A.R. Actin and α-actinin orchestrate the assembly and maturation of nascent adhesions in a myosin II motor-independent manner. Nat. Cell Biol. 2008;9:1039-1050.
Not in our library10.1038/ncb1763
354.
Ridley A.J. Life at the Leading Edge. Cell. 2011;145:1012-1022.
Not in our library10.1016/j.cell.2011.06.010
355.
Pasapera A.M., Plotnikov S.V., Fischer R.S., Case L.B., Egelhoff T.T., Waterman C.M. Rac1-Dependent Phosphorylation and Focal Adhesion Recruitment of Myosin IIA Regulates Migration and Mechanosensing. Curr. Biol. 2015;25:175-186.
Not in our library10.1016/j.cub.2014.11.043
356.
Kuo J.-C., Han X., Hsiao C.-T., Yates III J.R., Waterman C.M. Analysis of the myosin-II-responsive focal adhesion proteome reveals a role for β-Pix in negative regulation of focal adhesion maturation. Nat. Cell Biol. 2011;13:383-393.
Not in our library10.1038/ncb2216
357.
Schiller H.B., Friedel C.C., Boulegue C., Fässler R. Quantitative proteomics of the integrin adhesome show a myosin II-dependent recruitment of LIM domain proteins. EMBO Rep. 2011;12:259-266.
Not in our library10.1038/embor.2011.5
358.
Peng J., Wallar B.J., Flanders A., Swiatek P.J., Alberts A.S. Disruption of the Diaphanous-related formin Drf1 gene encoding mDia1 reveals a role for Drf3 as an effector for Cdc42. Curr. Biol. 2003;13:534-545.
Not in our library10.1016/s0960-9822(03)00170-2
359.
Iskratsch T., Yu C.H., Mathur A., Liu S., Stévenin V., Dwyer J., Hone J., Ehler E., Sheetz M. FHOD1 is needed for directed forces and adhesion maturation during cell spreading and migration. Dev. Cell. 2013;27:545-559.
Not in our library10.1016/j.devcel.2013.11.003
360.
DeMali K.A., Barlow C.A., Burridge K. Recruitment of the Arp2/3 complex to vinculin: Coupling membrane protrusion to matrix adhesion. J. Cell Biol. 2002;159:881-891.
Not in our library10.1083/jcb.200206043
361.
Serrels B., Serrels A., Brunton V.G., Holt M., McLean G.W., Gray C.H., Jones G.E., Frame M.C. Focal adhesion kinase controls actin assembly via a FERM-mediated interaction with the Arp2/3 complex. Nat. Cell Biol. 2007;9:1046-1056.
Not in our library10.1038/ncb1626
362.
Suraneni P., Rubinstein B., Unruh J.R., Durnin M., Hanein D., Li R. The Arp2/3 complex is required for lamellipodia extension and directional fibroblast cell migration. J. Cell Biol. 2012;197:239-251.
Not in our library10.1083/jcb.201112113
363.
Wu C., Asokan S.B., Berginski M.E., Haynes E.M., Sharpless N.E., Griffith J.D., Gomez S.M., Bear J.E. Arp2/3 Is Critical for Lamellipodia and Response to Extracellular Matrix Cues but Is Dispensable for Chemotaxis. Cell. 2012;148:973-987.
Not in our library10.1016/j.cell.2011.12.034
364.
Li A., Ma Y., Yu X., Mort R.L., Lindsay C.R., Stevenson D., Strathdee D., Insall R.H., Chernoff J., Snapper S.B., et al. Rac1 Drives Melanoblast Organization during Mouse Development by Orchestrating Pseudopod- Driven Motility and Cell-Cycle Progression. Dev. Cell. 2011;21:722-734.
Not in our library10.1016/j.devcel.2011.07.008
365.
O’Neill P.R., Kalyanaraman V., Gautman N. Subcellular optogenetic activation of Cdc42 controls local and distal signaling to drive immune cell migration. Mol. Biol. Cell. 2016;27:1442-1450.
Not in our library10.1091/mbc.e15-12-0832
366.
Yang H.W., Collins S.R., Meyer T. Locally excitable Cdc42 signals steer cells during chemotaxis. Nat. Cell Biol. 2016;18:191-201.
Not in our library10.1038/ncb3292
367.
Harris K.P., Tepass U. Cdc42 and Vesicle Trafficking in Polarized Cells. Traffic. 2010;11:1272-1279.
Not in our library10.1111/j.1600-0854.2010.01102.x
368.
Vega F.M., Fruhwirth G., Ng T., Ridley A.J. RhoA and RhoC have distinct roles in migration and invasion by acting through different targets. J. Cell Biol. 2011;193:655-665.
Not in our library10.1083/jcb.201011038
369.
Petrie R.J., Yamada K.M. At the leading edge of three-dimensional cell migration. J. Cell Sci. 2012;125:5917-5926.
Not in our library10.1242/jcs.093732
370.
Paluch E.K., Aspalter I.M., Sixt M. Focal Adhesion–Independent Cell Migration. Annu. Rev. Cell Dev. Biol. 2016;32:469-490.
Not in our library10.1146/annurev-cellbio-111315-125341
371.
Maugis B., Brugues J., Nassoy P., Guillen N., Sens P., Amblard F. Dynamic instability of the intracellular pressure drives bleb-based motility. J. Cell Sci. 2010;123:3884-3892.
Not in our library10.1242/jcs.065672
372.
Bergert M., Chandradoss S.D., Desai R.A., Paluch E. Cell mechanics control rapid transitions between blebs and lamellipodia during migration. Proc. Natl. Acad. Sci. USA. 2012;109:14434-14439.
Not in our library10.1073/pnas.1207968109
373.
Weiser D.C., Row R.H., Kimelman D. Rho-regulated Myosin phosphatase establishes the level of protrusive activity required for cell movements during zebrafish gastrulation. Development. 2009;136:2375-2384.
Not in our library10.1242/dev.034892
374.
Sroka J., von Gunten M., Dunn G.A., Keller H.U. Phenotype modulation in non-adherent and adherent sublines of Walker carcinosarcoma cells: The role of cell-substratum contacts and microtubules in controlling cell shape, locomotion and cytoskeletal structure. Int. J. Biochem. Cell Biol. 2002;34:882-899.
Not in our library10.1016/s1357-2725(01)00178-9
375.
Blaser H., Eisenbeiss S., Neumann M., Reichman-Fried M., Thisse B., Thisse C., Raz E. Transition from non-motile behaviour to directed migration during early PGC development in zebrafish. J. Cell Sci. 2005;118:4027-4038.
Not in our library10.1242/jcs.02522
376.
Malawista S.E., de Boisfleury Chevance A., Boxer L.A. Random locomotion and chemotaxis of human blood polymorphonuclear leukocytes from a patient with Leukocyte Adhesion Deficiency-1: Normal displacement in close quarters via chimneying. Cell Motil. Cytoskeleton. 2000;46:183-189.
Not in our library10.1002/1097-0169(200007)46:3<183::aid-cm3>3.0.co;2-2
377.
Tozluoǧlu M., Tournier A.L., Jenkins R.P., Hooper S., Bates P.A., Sahai E. Matrix geometry determines optimal cancer cell migration strategy and modulates response to interventions. Nat. Cell Biol. 2013;15:751-762.
Not in our library10.1038/ncb2775
378.
Macrophage-Induced Blood Vessels Guide Schwann Cell-Mediated Regeneration of Peripheral Nerves · 2015
No notice
379.
Martinelli S., Chen E.J.H., Clarke F., Lyck R., Affentranger S., Burkhardt J.K., Niggli V. Ezrin/Radixin/Moesin Proteins and Flotillins Cooperate to Promote Uropod Formation in T Cells. Front. Immunol. 2013;4:84.
Not in our library10.3389/fimmu.2013.00084
380.
Niggli V., Rossy J. Ezrin/radixin/moesin: Versatile controllers of signaling molecules and of the cortical cytoskeleton. Int. J. Biochem. Cell Biol. 2008;40:344-349.
Not in our library10.1016/j.biocel.2007.02.012
381.
Lorentzen A., Bamber J., Sadok A., Elson-Schwab I., Marshall C.J. An ezrin-rich, rigid uropod-like structure directs movement of amoeboid blebbing cells. J. Cell Sci. 2011;124:1256-1267.
Not in our library10.1242/jcs.074849
382.
Yanase Y., Hide I., Mihara S., Shirai Y., Saito N., Nakata Y., Hide M., Sakai N. A critical role of conventional protein kinase C in morphological changes of rodent mast cells. Immunol. Cell Biol. 2011;89:149-159.
Not in our library10.1038/icb.2010.67
383.
Fehon R.G., McClatchey A.I., Bretscher A. Organizing the cell cortex: The role of ERM proteins. Nat. Rev. Mol. Cell Biol. 2010;11:276-287.
Not in our library10.1038/nrm2866
384.
Evans E.A., Skalak R. Mechanics and Thermodynamics of Biomembranes: Part 1. CRC Crit. Rev. Bioeng. 1979;3:181-330.
Not in our library
385.
Pinner S., Sahai E. PDK1 regulates cancer cell motility by antagonising inhibition of ROCK1 by RhoE. Nat. Cell Biol. 2008;10:127-137.
Not in our library10.1038/ncb1675
386.
Trinkaus J.P. Ingression during Early Gastrulation of Fundulus. Dev. Biol. 1996;177:356-370.
Not in our library10.1006/dbio.1996.0169
387.
Oakes P.W., Beckham Y., Stricker J., Gardel M.L. Tension is required but not sufficient for focal adhesion maturation without a stress fiber template. J. Cell Biol. 2012;196:363-374.
Not in our library10.1083/jcb.201107042
388.
Puklin-Faucher E., Gao M., Schulten K., Vogel V. How the headpiece hinge angle is opened: New insights into the dynamics of integrin activation. J. Cell Biol. 2006;175:349-360.
Not in our library10.1083/jcb.200602071
389.
Gingras A.R., Vogel K.-P., Steinhoff H.-J., Zoegler W.H., Patel B., Emsley J., Critchley D.R., Roberts G.C.K., Barsukov I.L. Structural and Dynamic Characterization of a Vinculin Binding Site in the Talin Rod. Biochemistry. 2006.
Not in our library10.1021/bi052136l
390.
del Rio A., Perez-Jimenez R., Liu R., Roca-Cusachs P., Fernandez J.M., Sheetz M.P. Stretching Single Talin Rod Molecules Activates Vinculin Binding. Science. 2009;323:638-641.
Not in our library10.1126/science.1162912
391.
Johnson C.P., Tang H.-Y., Carag C., Speicher D.W., Discher D.E. Forced unfolding of proteins within cells. Science. 2007;317:663-666.
Not in our library10.1126/science.1139857
392.
Actin stress fibre subtypes in mesenchymal-migrating cells · 2013
No notice
393.
Valerius N.H., Stendahl O., Hartwig J.H., Stossel T.P. Distribution of actin-binding protein and myosin in polymorphonuclear leukocytes during locomotion and phagocytosis. Cell. 1981;24:195-202.
Not in our library10.1016/0092-8674(81)90515-8
394.
Rubino S., Fighetti M., Unger E., Cappuccinelli P. Location of actin, myosin, and microtubular structures during directed locomotion of Dictyostelium amebae. J. Cell Biol. 1984;98:382-390.
Not in our library10.1083/jcb.98.2.382
395.
Cheng S., Castillo V., Welty M., Eliaz I., Sliva D. Honokiol inhibits migration of renal cell carcinoma through activation of RhoA/ROCK/MLC signaling pathway. Int. J. Oncol. 2016;49:1525-1530.
Not in our library10.3892/ijo.2016.3663
396.
Castella L.F., Buscemi L., Godbout C., Meister J.-J., Hinz B. A new lock-step mechanism of matrix remodelling based on subcellular contractile events. J. Cell Sci. 2010;123:1751-1760.
Not in our library10.1242/jcs.066795
397.
Kirfel G., Rigort A., Borm B., Herzog V. Cell migration: Mechanisms of rear detachment and the formation of migration tracks. Eur. J. Cell Biol. 2004;83:717-724.
Not in our library10.1078/0171-9335-00421
398.
Vicente-Manzanares M., Koach M.A., Whitmore L., Lamers M.L., Horwitz A.F. Segregation and activation of myosin IIB creates a rear in migrating cells. J. Cell Biol. 2008;183:543-554.
Not in our library10.1083/jcb.200806030
399.
Cuddihy A.R., Bristow R.G. The p53 protein family and radiation sensitivity: Yes or no? Cancer Metastasis Rev. 2004;23:237-257.
Not in our library10.1023/b:canc.0000031764.81141.e4
400.
Totsukawa G., Wu Y., Sasaki Y., Hartshorne D.J., Yamakita Y., Yamashiro S., Matsumura F. Distinct roles of MLCK and ROCK in the regulation of membrane protrusions and focal adhesion dynamics during cell migration of fibroblasts. J. Cell Biol. 2004;164:427-439.
Not in our library10.1083/jcb.200306172
401.
Aifuwa I., Giri A., Longe N., Lee S.H., An S.S., Wirtz D. Senescent stromal cells induce cancer cell migration via inhibition of RhoA/ROCK/myosin-based cell contractility. Oncotarget. 2015;6:30516-30531.
Not in our library10.18632/oncotarget.5854
402.
Schmidt C.E., Horwitz A.F., Lauffenburger D.A., Sheetz M.P. Integrin-cytoskeletal interactions in migrating fibroblasts are dynamic, asymmetric, and regulated. J. Cell Biol. 1993;123:977-991.
Not in our library10.1083/jcb.123.4.977
403.
Brangwynne C.P., MacKintosh F.C., Weitz D.A. Force fluctuations and polymerization dynamics of intracellular microtubules. Proc. Natl. Acad. Sci. USA. 2007;104:16128-16133.
Not in our library10.1073/pnas.0703094104
404.
Wadsworth P. Regional regulation of microtubule dynamics in polarized, motile cells. Cell Motil. Cytoskeleton. 1999;42:48-59.
Not in our library10.1002/(sici)1097-0169(1999)42:1<48::aid-cm5>3.0.co;2-8
405.
Schober J.M., Cain J.M., Komarova Y.A., Borisy G.G. Migration and actin protrusion in melanoma cells are regulated by EB1 protein. Cancer Lett. 2009;284:30-36.
Not in our library10.1016/j.canlet.2009.04.007
406.
Al-Bassam J., Kim H., Brouhard G., van Oijen A., Harrison S.C., Chang F. CLASP Promotes Microtubule Rescue by Recruiting Tubulin Dimers to the Microtubule. Dev. Cell. 2010;19:245-258.
Not in our library10.1016/j.devcel.2010.07.016
407.
Steinmetz M.O., Jahnke W., Towbin H., Garcia-Echeverria C., Voshol H., Muller D., van Oostrum J. Phosphorylation disrupts the central helix in Op18/stathmin and suppresses binding to tubulin. EMBO Rep. 2001;2:505-510.
Not in our library10.1093/embo-reports/kve105
408.
Single Molecule Imaging Reveals Differences in Microtubule Track Selection Between Kinesin Motors · 2009
No notice
409.
Fygenson D.K., Marko J.F., Libchaber A. Mechanics of microtubule-based membrane extension. Phys. Rev. Lett. 1997;79:4497-4500.
Not in our library10.1103/physrevlett.79.4497
410.
Mogilner A., Oster G. Polymer motors: Pushing out the front and pulling up the back. Curr. Biol. 2003;13:721-733.
Not in our library10.1016/j.cub.2003.08.050
411.
Yu W., Centonze V.E., Ahmad F.J., Baas P.W. Microtubule nucleation and release from the neuronal centrosome. J. Cell Biol. 1993;122:349-359.
Not in our library10.1083/jcb.122.2.349
412.
Roll-Mecak A., McNally F.J. Microtubule-severing enzymes. Curr. Opin. Cell Biol. 2010;22:96-103.
Not in our library10.1016/j.ceb.2009.11.001
413.
Marcette J.D., Chen J.J., Nonet M.L. The Caenorhabditis elegans microtubule minus-end binding homolog PTRN-1 stabilizes synapses and neurites. Elife. 2014;3:e01637.
Not in our library10.7554/elife.01637
414.
Richardson C.E., Spilker K.A., Cueva J.G., Perrino J., Goodman M.B., Shen K. PTRN-1, a microtubule minus end-binding CAMSAP homolog, promotes microtubule function in Caenorhabditis elegans neurons. Elife. 2014;3:e01498.
Not in our library10.7554/elife.01498
415.
Yau K.W., Schatzle P., Tortosa E., Pages S., Holtmaat A., Kapitein L.C., Hoogenraad C.C. Dendrites In Vitro and In Vivo Contain Microtubules of Opposite Polarity and Axon Formation Correlates with Uniform Plus-End-Out Microtubule Orientation. J. Neurosci. 2016;36:1071-1085.
Not in our library10.1523/jneurosci.2430-15.2016
416.
Baas P.W., Deitch J.S., Black M.M., Banker G.A. Polarity orientation of microtubules in hippocampal neurons: Uniformity in the axon and nonuniformity in the dendrite. Proc. Natl. Acad. Sci. USA. 1988;85:8335-8339.
Not in our library10.1073/pnas.85.21.8335
417.
Bradke F., Dotti C.G. The role of local actin instability in axon formation. Science. 1999;283:1931-1934.
Not in our library10.1126/science.283.5409.1931
418.
Wiesner C., Faix J., Himmel M., Bentzien F., Linder S. KIF5B and KIF3A/KIF3B kinesins drive MT1-MMP surface exposure, CD44 shedding, and extracellular matrix degradation in primary macrophages. Blood. 2010;116:1559-1569.
Not in our library10.1182/blood-2009-12-257089
419.
Steffen A., Le Dez G., Poincloux R., Recchi C., Nassoy P., Rottner K., Galli T., Chavrier P. MT1-MMP-Dependent Invasion Is Regulated by TI-VAMP/VAMP7. Curr. Biol. 2008;18:926-931.
Not in our library10.1016/j.cub.2008.05.044
420.
Talin-KANK1 interaction controls the recruitment of cortical microtubule stabilizing complexes to focal adhesions · 2016
No notice
421.
Stehbens S., Wittmann T. Targeting and transport: How microtubules control focal adhesion dynamics. J. Cell Biol. 2012;198:481-489.
Not in our library10.1083/jcb.201206050
422.
Rooney C., White G., Nazgiewicz A., Woodcock S.A., Anderson K.I., Ballestrem C., Malliri A. The Rac activator STEF (Tiam2) regulates cell migration by microtubule-mediated focal adhesion disassembly. EMBO Rep. 2010;11:292-298.
Not in our library10.1038/embor.2010.10
423.
Rodriguez O.C., Schaefer A.W., Mandato C.A., Forscher P., Bement W.M., Waterman-Storer C.M. Conserved microtubule–actin interactions in cell movement and morphogenesis. Nat. Cell Biol. 2003;5:599-609.
Not in our library10.1038/ncb0703-599
424.
Krendel M., Zenke F.T., Bokoch G.M. Nucleotide exchange factor GEF-H1 mediates cross-talk between microtubules and the actin cytoskeleton. Nat. Cell Biol. 2002;4:294-301.
Not in our library10.1038/ncb773
425.
Waterman-Storer C.M., Worthylake R.A., Liu B.P., Burridge K., Salmon E.D. Microtubule growth activates Rac1 to promote lamellipodial protrusion in fibroblasts. Nat. Cell Biol. 1999;1:45-50.
Not in our library10.1038/9018
426.
Montenegro-Venegas C., Tortosa E., Rosso S., Peretti D., Bollati F., Bisbal M., Jausoro I., Avila J., Cáceres A., Gonzalez-Billault C. MAP1B regulates axonal development by modulating Rho-GTPase Rac1 activity. Mol. Biol. Cell. 2010;21:3518-3528.
Not in our library10.1091/mbc.e09-08-0709
427.
Van Haren J., Boudeau J., Schmidt S., Basu S., Liu Z., Lammers D., Demmers J., Benhari J., Grosveld F., Debant A., et al. Dynamic microtubules catalyze formation of navigator-TRIO complexes to regulate neurite extension. Curr. Biol. 2014;24:1778-1785.
Not in our library10.1016/j.cub.2014.06.037
428.
Nalbant P., Chang Y.-C., Birkenfeld J., Chang Z.-F., Bokoch G.M. Guanine nucleotide exchange factor-H1 regulates cell migration via localized activation of RhoA at the leading edge. Mol. Biol. Cell. 2009;20:4070-4082.
Not in our library10.1091/mbc.e09-01-0041
429.
Callow M.G., Zozulya S., Gishizky M.L., Jallal B., Smeal T. PAK4 mediates morphological changes through the regulation of GEF-H1. J. Cell Sci. 2005;118:1861-1872.
Not in our library10.1242/jcs.02313
430.
Ren Y., Li R., Zheng Y., Busch H. Cloning and characterization of GEF-H1, a microtubule-associated guanine nucleotide exchange factor for Rac and Rho GTPases. J. Biol. Chem. 1998;273:34954-34960.
Not in our library10.1074/jbc.273.52.34954
431.
Zenke F.T., Krendel M., DerMardirossian C., King C.C., Bohl B.P., Bokoch G.M. p21-activated Kinase 1 Phosphorylates and Regulates 14-3-3 Binding to GEF-H1, a Microtubule-localized Rho Exchange Factor. J. Biol. Chem. 2004;279:18392-18400.
Not in our library10.1074/jbc.m400084200
432.
Kaverina I., Rottner K., Small J.V. Targeting, capture, and stabilization of microtubules at early focal adhesions. J. Cell Biol. 1998;142:181-190.
Not in our library10.1083/jcb.142.1.181
433.
Gu Z., Noss E.H., Hsu V.W., Brenner M.B. Integrins traffic rapidly via circular dorsal ruffles and macropinocytosis during stimulated cell migration. J. Cell Biol. 2011;193:61-70.
Not in our library10.1083/jcb.201007003
434.
López M.P., Huber F., Grigoriev I., Steinmetz M.O., Akhmanova A., Koenderink G.H., Dogterom M. Actin–microtubule coordination at growing microtubule ends. Nat. Commun. 2014;5:4778.
Not in our library10.1038/ncomms5778
435.
Wickström S.A., Fässler R. Regulation of membrane traffic by integrin signaling. Trends Cell Biol. 2011;21:266-273.
Not in our library10.1016/j.tcb.2011.02.003
436.
Etienne-Manneville S. APC in cell migration. Adv. Exp. Med. Biol. 2009;656:30-40.
Not in our library10.1007/978-1-4419-1145-2_3
437.
Matsumoto S., Fumoto K., Okamoto T., Kaibuchi K., Kikuchi A. Binding of APC and dishevelled mediates Wnt5a-regulated focal adhesion dynamics in migrating cells. EMBO J. 2010;29:1192-1204.
Not in our library10.1038/emboj.2010.26
438.
Juanes M.A., Bouguenina H., Eskin J.A., Jaiswal R., Badache A., Goode B.L. Adenomatous polyposis coli nucleates actin assembly to drive cell migration and microtubuleinduced focal adhesion turnover. J. Cell Biol. 2017;216:2859-2875.
Not in our library10.1083/jcb.201702007
439.
Breitsprecher D., Jaiswal R., Bombardier J.P., Gould C.J., Gelles J., Goode B.L. Rocket Launcher Mechanism of Collaborative Actin Assembly Defined by Single-Molecule Imaging. Science. 2012;336:1164-1168.
Not in our library10.1126/science.1218062
440.
Brandt D.T., Grosse R. Get to grips: Steering local actin dynamics with IQGAPs. EMBO Rep. 2007;8:1019-1023.
Not in our library10.1038/sj.embor.7401089
441.
Krylyshkina O., Anderson K.I., Kaverina I., Upmann I., Manstein D.J., Small J.V., Toomre D.K. Nanometer targeting of microtubules to focal adhesions. J. Cell Biol. 2003;161:853-859.
Not in our library10.1083/jcb.200301102
442.
Kaverina I., Krylyshkina O., Small J.V. Microtubule targeting of substrate contacts promotes their relaxation and dissociation. J. Cell Biol. 1999;146:1033-1043.
Not in our library10.1083/jcb.146.5.1033
443.
Rid R., Schiefermeier N., Grigoriev I., Small J.V., Kaverina I. The last but not the least: The origin and significance of trailing adhesions in fibroblastic cells. Cell Motil. Cytoskeleton. 2005;61:161-171.
Not in our library10.1002/cm.20076
444.
Applewhite D.A., Grode K.D., Keller D., Zadeh A.D., Zadeh A., Slep K.C., Rogers S.L. The spectraplakin Short stop is an actin-microtubule cross-linker that contributes to organization of the microtubule network. Mol. Biol. Cell. 2010;21:1714-1724.
Not in our library10.1091/mbc.e10-01-0011
445.
Wu X., Shen Q.-T., Oristian D.S., Lu C.P., Zheng Q., Wang H.-W., Fuchs E. Skin stem cells orchestrate directional migration by regulating microtubule-ACF7 connections through GSK3β. Cell. 2011;144:341-352.
Not in our library10.1016/j.cell.2010.12.033
446.
Zaoui K., Benseddik K., Daou P., Salaun D., Badache A. ErbB2 receptor controls microtubule capture by recruiting ACF7 to the plasma membrane of migrating cells. Proc. Natl. Acad. Sci. USA. 2010;107:18517-18522.
Not in our library10.1073/pnas.1000975107
447.
Higashida C., Miyoshi T., Fujita A., Oceguera-Yanez F., Monypenny J., Andou Y., Narumiya S., Watanabe N. Actin Polymerization-Driven Molecular Movement of mDia1 in Living Cells. Science. 2004;303:2007-2010.
Not in our library10.1126/science.1093923
448.
Wickström S.A., Lange A., Hess M.W., Polleux J., Spatz J.P., Krüger M., Pfaller K., Lambacher A., Bloch W., Mann M., et al. Integrin-Linked Kinase Controls Microtubule Dynamics Required for Plasma Membrane Targeting of Caveolae. Dev. Cell. 2010;19:574-588.
Not in our library10.1016/j.devcel.2010.09.007
449.
Villari G., Jayo A., Zanet J., Fitch B., Serrels B., Frame M., Stramer B., Goult B., Parsons M. A direct interaction between fascin and microtubules contributes to adhesion dynamics and cell migration. J. Cell Sci. 2015;128:4601-4604.
Not in our library10.1242/jcs.175760
450.
Ezratty E.J., Partridge M.A., Gundersen G.G. Microtubule-induced focal adhesion disassembly is mediated by dynamin and focal adhesion kinase. Nat. Cell Biol. 2005;7:581-590.
Not in our library10.1038/ncb1262
451.
Ezratty E.J., Bertaux C., Marcantonio E.E., Gundersen G.G. Clathrin mediates integrin endocytosis for focal adhesion disassembly in migrating cells. J. Cell Biol. 2009;187:733-747.
Not in our library10.1083/jcb.200904054
452.
Stehbens S.J., Paszek M., Pemble H., Ettinger A., Gierke S., Wittmann T. CLASPs link focal-adhesion-associated microtubule capture to localized exocytosis and adhesion site turnover. Nat. Cell Biol. 2014;16:558-570.
Not in our library10.1038/ncb2975
453.
Kenific C.M., Stehbens S.J., Goldsmith J., Leidal A.M., Faure N., Ye J., Wittmann T., Debnath J. NBR 1 enables autophagy-dependent focal adhesion turnover. J. Cell Biol. 2016;212:577-590.
Not in our library10.1083/jcb.201503075
454.
Helfand B.T., Mendez M.G., Murthy S.N.P., Shumaker D.K., Grin B., Mahammad S., Aebi U., Wedig T., Wu Y.I., Hahn K.M., et al. Vimentin organization modulates the formation of lamellipodia. Mol. Biol. Cell. 2011;22:1274-1289.
Not in our library10.1091/mbc.e10-08-0699
455.
Menko A.S., Bleaken B.M., Libowitz A.A., Zhang L., Stepp M.A., Walker J.L. A central role for vimentin in regulating repair function during healing of the lens epithelium. Mol. Biol. Cell. 2014;25:776-790.
Not in our library10.1091/mbc.e12-12-0900
456.
Sakamoto Y., Boëda B., Etienne-Manneville S. APC binds intermediate filaments and is required for their reorganization during cell migration. J. Cell Biol. 2013;200:249-258.
Not in our library10.1083/jcb.201206010
457.
Huber F., Boire A., López M.P., Koenderink G.H. Cytoskeletal crosstalk: When three different personalities team up. Curr. Opin. Cell Biol. 2015;32:39-47.
Not in our library10.1016/j.ceb.2014.10.005
458.
Shabbir S.H., Cleland M.M., Goldman R.D., Mrksich M. Geometric control of vimentin intermediate filaments. Biomaterials. 2014;35:1359-1366.
Not in our library10.1016/j.biomaterials.2013.10.008
459.
Yuan A., Rao M.V., Veeranna, Nixon R.A. Neurofilaments and neurofilament proteins in health and disease. Cold Spring Harb. Perspect. Biol. 2017;9.
Not in our library10.1101/cshperspect.a018309
460.
Mendez M.G., Kojima S.-I., Goldman R.D. Vimentin induces changes in cell shape, motility, and adhesion during the epithelial to mesenchymal transition. FASEB J. 2010;24:1838-1851.
Not in our library10.1096/fj.09-151639
461.
Rogel M.R., Soni P.N., Troken J.R., Sitikov A., Trejo H.E., Ridge K.M. Vimentin is sufficient and required for wound repair and remodeling in alveolar epithelial cells. FASEB J. 2011;25:3873-3883.
Not in our library10.1096/fj.10-170795
462.
Chung B.M., Rotty J.D., Coulombe P.A. Networking galore: Intermediate filaments and cell migration. Curr. Opin. Cell Biol. 2013;25:600-612.
Not in our library10.1016/j.ceb.2013.06.008
463.
Vimentin on the move: new developments in cell migration · 2018
No notice
464.
Lepekhin E.A., Eliasson C., Berthold C.H., Berezin V., Bock E., Pekny M. Intermediate filaments regulate astrocyte motility. J. Neurochem. 2001;79:617-625.
Not in our library10.1046/j.1471-4159.2001.00595.x
465.
Rathje L.-S.Z., Nordgren N., Pettersson T., Ronnlund D., Widengren J., Aspenstrom P., Gad A.K.B. Oncogenes induce a vimentin filament collapse mediated by HDAC6 that is linked to cell stiffness. Proc. Natl. Acad. Sci. USA. 2014;111:1515-1520.
Not in our library10.1073/pnas.1300238111
466.
Gan Z., Ding L., Burckhardt C.J., Lowery J., Zaritsky A., Sitterley K., Mota A., Costigliola N., Starker C.G., Voytas D.F., et al. Vimentin Intermediate Filaments Template Microtubule Networks to Enhance Persistence in Cell Polarity and Directed Migration. Cell Syst. 2016;3:252-263.
Not in our library10.1016/j.cels.2016.08.007
467.
The Roles of Post-translational Modifications in the Context of Protein Interaction Networks · 2015
No notice
468.
Nieuwenhuizen R.P.J., Nahidiazar L., Manders E.M.M., Jalink K., Stallinga S., Rieger B. Co-orientation: Quantifying simultaneous co-localization and orientational alignment of filaments in light microscopy. PLoS ONE. 2015;10:1-21.
Not in our library10.1371/journal.pone.0131756
469.
Leduc C., Manneville S.E. Regulation of microtubule-associated intermediate filament network polarization. J. Cell Biol. 2017;216:1689-1703.
Not in our library10.1083/jcb.201607045
470.
Havel L., Kline E., Salgueiro A., Marcus A. Vimentin regulates lung cancer cell adhesion through a VAV2–Rac1 pathway to control focal adhesion kinase activity. Oncogene. 2014;34:1-12.
Not in our library10.1038/onc.2014.123
471.
Vimentin intermediate filaments control actin stress fiber assembly through GEF-H1 and RhoA · 2017
No notice
472.
De Pascalis C., Perez-Gonzalez C., Seetharaman S., Boeda B., Vianay B., Burute M., Leduc C., Borghi N., Trepat X., Etienne-Manneville S. Intermediate filaments control collective migration by restricting traction forces and sustaining cell-cell contacts. bioRxiv. 2018.
Not in our library10.1083/jcb.201801162
473.
Kim J., Yang C., Kim E.J., Jang J., Kim S.-J., Kang S.M., Kim M.G., Jung H., Park D., Kim C. Vimentin filaments regulate integrin–ligand interactions by binding to the cytoplasmic tail of integrin β3. J. Cell Sci. 2016;129:2030-2042.
Not in our library10.1242/jcs.180315
474.
Costigliola N., Ding L., Burckhardt C.J., Han S.J., Gutierrez E., Mota A., Groisman A., Mitchison T.J., Danuser G. Vimentin fibers orient traction stress. Proc. Natl. Acad. Sci. USA. 2017;114:5195-5200.
Not in our library10.1073/pnas.1614610114
475.
Akhmanova A., Stehbens S.J., Yap A.S. Touch, Grasp, Deliver and Control: Functional Cross-Talk Between Microtubules and Cell Adhesions. Traffic. 2009;10:268-274.
Not in our library10.1111/j.1600-0854.2008.00869.x
476.
Janosch P., Kieser A., Eulitz M., Lovric J., Sauer G., Reichert M., Gounari F., Büscher D., Baccarini M., Mischak H., et al. The Raf-1 kinase associates with vimentin kinases and regulates the structure of vimentin filaments. FASEB J. 2000;14:2008-2021.
Not in our library10.1096/fj.99-0883com
477.
Ehrenreiter K., Piazzolla D., Velamoor V., Sobczak I., Small J.V., Takeda J., Leung T., Baccarini M. Raf-1 regulates Rho signaling and cell migration. J. Cell Biol. 2005;168:955-964.
Not in our library10.1083/jcb.200409162
478.
Amano M., Fukata Y., Kaibuchi K. Regulation and Functions of Rho-Associated Kinase. Exp. Cell Res. 2000;261:44-51.
Not in our library10.1006/excr.2000.5046
479.
Sin W.C., Chen X.Q., Leung T., Lim L. RhoA-binding kinase alpha translocation is facilitated by the collapse of the vimentin intermediate filament network. Mol. Cell. Biol. 1998;18:6325-6339.
Not in our library10.1128/mcb.18.11.6325
480.
Ren X.D., Kiosses W.B., Sieg D.J., Otey C.A., Schlaepfer D.D., Schwartz M.A. Focal adhesion kinase suppresses Rho activity to promote focal adhesion turnover. J. Cell Sci. 2000;113:3673-3678.
Not in our library10.1242/jcs.113.20.3673
481.
Ren X.D., Kiosses W.B., Schwartz M.A. Regulation of the small GTP-binding protein Rho by cell adhesion and the cytoskeleton. EMBO J. 1999;18:578-585.
Not in our library10.1093/emboj/18.3.578
482.
Jiu Y., Lehtimäki J., Tojkander S., Cheng F., Jäälinoja H., Liu X., Varjosalo M., Eriksson J.E., Lappalainen P. Bidirectional Interplay between Vimentin Intermediate Filaments and Contractile Actin Stress Fibers. Cell Rep. 2015;11:1511-1518.
Not in our library10.1016/j.celrep.2015.05.008
483.
Goto H., Tanabe K., Manser E., Lim L., Yasui Y., Inagaki M. Phosphorylation and reorganization of vimentin by p21-activated kinase (PAK). Genes Cells. 2002;7:91-97.
Not in our library10.1046/j.1356-9597.2001.00504.x
484.
Petrie R.J., Koo H., Yamada K.M. Generation of compartmentalized pressure by a nuclear piston governs cell motility in a 3D matrix. Science. 2014;345:1062-1065.
Not in our library10.1126/science.1256965
485.
Sun N., Huiatt T.W., Paulin D., Li Z., Robson R.M. Synemin interacts with the LIM domain protein zyxin and is essential for cell adhesion and migration. Exp. Cell Res. 2010;316:491-505.
Not in our library10.1016/j.yexcr.2009.10.015
486.
Pan Y., Jing R., Pitre A., Williams B.J., Skalli O. Intermediate filament protein synemin contributes to the migratory properties of astrocytoma cells by influencing the dynamics of the actin cytoskeleton. FASEB J. 2008;22:3196-3206.
Not in our library10.1096/fj.08-106187
487.
Uyama N., Zhao L., Van Rossen E., Hirako Y., Reynaert H., Adams D.H., Xue Z., Li Z., Robson R., Pekny M., et al. Hepatic stellate cells express synemin, a protein bridging intermediate filaments to focal adhesions. Gut. 2006;55:1276-1289.
Not in our library10.1136/gut.2005.078865
488.
Sun N., Critchley D.R., Paulin D., Li Z., Robson R.M. Identification of a repeated domain within mammalian α-synemin that interacts directly with talin. Exp. Cell Res. 2008;314:1839-1849.
Not in our library10.1016/j.yexcr.2008.01.034
489.
Sun N., Critchley D.R., Paulin D., Li Z., Robson R.M. Human alpha-synemin interacts directly with vinculin and metavinculin. Biochem. J. 2008;409:657-667.
Not in our library10.1042/bj20071188
490.
Hijikata T., Nakamura A., Isokawa K., Imamura M., Yuasa K., Ishikawa R., Kohama K., Takeda S., Yorifuji H. Plectin 1 links intermediate filaments to costameric sarcolemma through -synemin, -dystrobrevin and actin. J. Cell Sci. 2008;121:2062-2074.
Not in our library10.1242/jcs.021634
491.
Hyder C.L., Lazaro G., Pylvänäinen J.W., Roberts M.W.G., Qvarnström S.M., Eriksson J.E. Nestin regulates prostate cancer cell invasion by influencing the localisation and functions of FAK and integrins. J. Cell Sci. 2014;127:2161-2173.
Not in our library10.1242/jcs.125062
492.
Projections of Global Mortality and Burden of Disease from 2002 to 2030 · 2006
No notice
493.
Ferlay J., Shin H.R., Bray F., Forman D., Mathers C., Parkin D.M. Estimates of worldwide burden of cancer in 2008: Globocan 2008. Int. J. Cancer. 2010;127:2893-2917.
Not in our library10.1002/ijc.25516
494.
Legler J.M., Ries L.A., Smith M.A., Warren J.L., Heineman E.F., Kaplan R.S., Linet M.S. Cancer surveillance series [corrected]: Brain and other central nervous system cancers: Recent trends in incidence and mortality. J. Natl. Cancer Inst. 1999;91:1382-1390.
Not in our library10.1093/jnci/91.16.1382
495.
Davis F.G., McCarthy B.J., Freels S., Kupelian V., Bondy M.L. The conditional probability of survival of patients with primary malignant brain tumors: Surveillance, epidemiology, and end results (SEER) data. Cancer. 1999;85:485-491.
Not in our library10.1002/(sici)1097-0142(19990115)85:2<485::aid-cncr29>3.0.co;2-l
496.
Scherer H.J. The forms of growth in gliomas and their practical significance. Brain. 1940;63:1-35.
Not in our library10.1093/brain/63.1.1
497.
Maher E.A., Furnari F.B., Bachoo R.M., Rowitch D.H., Louis D.N., Cavenee W.K., DePinho R.A. Malignant glioma: Genetics and biology of a grave matter. Genes Dev. 2001;15:1311-1333.
Not in our library10.1101/gad.891601
498.
Chamberlain M.C. Radiographic patterns of relapse in glioblastoma. J. Neurooncol. 2011;101:319-323.
Not in our library10.1007/s11060-010-0251-4
499.
Iwadate Y., Sakaida T., Saegusa T., Hiwasa T., Takiguchi M., Fujimoto S., Yamaura A. Proteome-based identification of molecular markers predicting chemosensitivity to each category of anticancer agents in human gliomas. Int. J. Oncol. 2005;26:993-998.
Not in our library10.3892/ijo.26.4.993
500.
Hirata E., Yukinaga H., Kamioka Y., Arakawa Y., Miyamoto S., Okada T., Sahai E., Matsuda M. In vivo fluorescence resonance energy transfer imaging reveals differential activation of Rho-family GTPases in glioblastoma cell invasion. J. Cell Sci. 2012;125:858-868.
Not in our library10.1242/jcs.089995
501.
Salhia B., Tran N.L., Chan A., Wolf A., Nakada M., Rutka F., Ennis M., McDonough W.S., Berens M.E., Symons M., et al. The Guanine Nucleotide Exchange Factors Trio, Ect2, and Vav3 Mediate the Invasive Behavior of Glioblastoma. Am. J. Pathol. 2008;173:1828-1838.
Not in our library10.2353/ajpath.2008.080043
502.
Malchinkhuu E., Sato K., Maehama T., Mogi C., Tomura H., Ishiuchi S., Yoshimoto Y., Kurose H., Okajima F. S1P2 receptors mediate inhibition of glioma cell migration through Rho signaling pathways independent of PTEN. Biochem. Biophys. Res. Commun. 2008;366:963-968.
Not in our library10.1016/j.bbrc.2007.12.054
503.
Tran N.L., McDonough W.S., Savitch B.A., Fortin S.P., Winkles J.A., Symons M., Nakada M., Cunliffe H.E., Hostetter G., Hoelzinger D.B., et al. Increased Fibroblast Growth Factor-Inducible 14 Expression Levels Promote Glioma Cell Invasion via Rac1 and Nuclear Factor-κB and Correlate with Poor Patient Outcome. Cancer Res. 2006;66:9535-9542.
Not in our library10.1158/0008-5472.can-06-0418
504.
Zagzag D., Friedlander D.R., Margolis B., Grumet M., Semenza G.L., Zhong H., Simons J.W., Holash J., Wiegand S.J., Yancopoulos G.D. Molecular Events Implicated in Brain Tumor Angiogenesis and Invasion. Pediatr. Neurosurg. 2000;33:49-55.
Not in our library10.1159/000028975
505.
Khalil B.D., Hanna S., Saykali B.A., El-Sitt S., Nasrallah A., Marston D., El-Sabban M., Hahn K.M., Symons M., El-Sibai M. The regulation of RhoA at focal adhesions by StarD13 is important for astrocytoma cell motility. Exp. Cell Res. 2014;321:109-122.
Not in our library10.1016/j.yexcr.2013.11.023
506.
Seasholtz T.M., Radeff-Huang J., Sagi S.A., Matteo R., Weems J.M., Cohen A.S., Feramisco J.R., Brown J.H. Rho-mediated cytoskeletal rearrangement in response to LPA is functionally antagonized by Rac1 and PIP2. J. Neurochem. 2004;91:501-512.
Not in our library10.1111/j.1471-4159.2004.02749.x
507.
Salhia B., Rutten F., Nakada M., Beaudry C., Berens M., Kwan A., Rutka J.T. Inhibition of Rho-kinase affects astrocytoma morphology, motility, and invasion through activation of Rac1. Cancer Res. 2005;65:8792-8800.
Not in our library10.1158/0008-5472.can-05-0160
508.
Chan A.Y., Coniglio S.J., Chuang Y., Michaelson D., Knaus U.G., Philips M.R., Symons M. Roles of the Rac1 and Rac3 GTPases in human tumor cell invasion. Oncogene. 2005;24:7821-7829.
Not in our library10.1038/sj.onc.1208909
509.
Chuang Y., Tran N.L., Rusk N., Nakada M., Berens M.E., Symons M. Role of Synaptojanin 2 in Glioma Cell Migration and Invasion. Cancer Res. 2004;64:8271-8275.
Not in our library10.1158/0008-5472.can-04-2097
510.
Yin M., Lu Q., Liu X., Wang T., Liu Y., Chen L. Silencing Drp1 inhibits glioma cells proliferation and invasion by RHOA/ROCK1 pathway. Biochem. Biophys. Res. Commun. 2016;478:663-668.
Not in our library10.1016/j.bbrc.2016.08.003
511.
Singh J., Sharma K., Pillai P.P. PDGFR inhibition mediated intracellular signalling in C6 glioma growth and migration: Role of ERK and ROCK pathway. Cytotechnology. 2018;70:465-477.
Not in our library10.1007/s10616-017-0163-7
512.
Kim D.-H., Wirtz D. Predicting how cells spread and migrate. Cell Adh. Migr. 2013;7:293-296.
Not in our library10.4161/cam.24804
513.
Fortin S.P., Ennis M.J., Schumacher C.A., Zylstra-Diegel C.R., Williams B.O., Ross J.T.D., Winkles J.A., Loftus J.C., Symons M.H., Tran N.L. Cdc42 and the Guanine Nucleotide Exchange Factors Ect2 and Trio Mediate Fn14-Induced Migration and Invasion of Glioblastoma Cells. Mol. Cancer Res. 2012;10:958-968.
Not in our library10.1158/1541-7786.mcr-11-0616
514.
Yamana N., Arakawa Y., Nishino T., Kurokawa K., Tanji M., Itoh R.E., Monypenny J., Ishizaki T., Bito H., Nozaki K., et al. The Rho-mDia1 Pathway Regulates Cell Polarity and Focal Adhesion Turnover in Migrating Cells through Mobilizing Apc and c-Src. Mol. Cell. Biol. 2006;26:6844-6858.
Not in our library10.1128/mcb.00283-06
515.
Ding L., Sun X., You Y., Liu N., Fu Z. Expression of focal adhesion kinase and phosphorylated focal adhesion kinase in human gliomas is associated with unfavorable overall survival. Transl. Res. 2010;156:45-52.
Not in our library10.1016/j.trsl.2010.05.001
516.
Jones G., Machado J., Merlo A. Loss of focal adhesion kinase (FAK) inhibits epidermal growth factor receptor-dependent migration and induces aggregation of nh(2)-terminal FAK in the nuclei of apoptotic glioblastoma cells. Cancer Res. 2001;61:4978-4981.
Not in our library
517.
Obara S., Nakata M., Takeshima H., Kuratsu J., Maruyama I., Kitajima I. Inhibition of migration of human glioblastoma cells by cerivastatin in association with focal adhesion kinase (FAK). Cancer Lett. 2002;185:153-161.
Not in our library10.1016/s0304-3835(02)00278-1
518.
Beadle C., Assanah M., Monzo P., Vallee R., Rosenfeld S., Canoll P. The Role of Myosin II in Glioma Invasion of the Brain Christopher. Mol. Biol. Cell. 2008;19:3357-3368.
Not in our library10.1091/mbc.e08-03-0319
519.
Ivkovic S., Beadle C., Noticewala S., Massey S.C., Swanson K.R., Toro L.N., Bresnick A.R., Canoll P., Rosenfeld S.S. Direct inhibition of myosin II effectively blocks glioma invasion in the presence of multiple motogens. Mol. Biol. Cell. 2012;23:533-542.
Not in our library10.1091/mbc.e11-01-0039
520.
Lee W., Lim S., Kim Y. The role of myosin II in glioma invasion: A mathematical model. PLoS ONE. 2017;12:1939-4586.
Not in our library10.1371/journal.pone.0171312
521.
Liu Z., Yang X., Chen C., Liu B., Ren B., Wang L., Zhao K., Yu S., Ming H. Expression of the Arp2/3 complex in human gliomas and its role in the migration and invasion of glioma cells. Oncol. Rep. 2013;30:2127-2136.
Not in our library10.3892/or.2013.2669
522.
Monzo P., Chong Y.K., Guetta-Terrier C., Krishnasamy A., Sathe S.R., Yim E.K.F., Ng W.H., Ang B.T., Tang C., Ladoux B., et al. Mechanical confinement triggers glioma linear migration dependent on formin FHOD3. Mol. Biol. Cell. 2016;27:1246-1261.
Not in our library10.1091/mbc.e15-08-0565
523.
Zhang C., Hai L., Zhu M., Yu S.P., Li T., Lin Y., Liu B., Zhou X.C., Chen L., Zhao P.F., et al. Actin cytoskeleton regulator Arp2/3 complex is required for DLL1 activating Notch1 signaling to maintain the stem cell phenotype of glioma initiating cells. Oncotarget. 2017;8:33353-33364.
Not in our library10.18632/oncotarget.16495
524.
Wang L., Zhao K., Ren B., Zhu M., Zhang C., Zhao P., Zhou H., Chen L., Yu S., Yang X. Expression of cortactin in human gliomas and its effect on migration and invasion of glioma cells. Oncol. Rep. 2015;34:1815-1824.
Not in our library10.3892/or.2015.4156
525.
Hwang J.H., Smith C.A., Salhia B., Rutka J.T. The Role of Fascin in the Migration and Invasiveness of Malignant Glioma Cells. Neoplasia. 2008;10:149-159.
Not in our library10.1593/neo.07909
526.
Hoa N.T., Ge L., Erickson K.L., Kruse C.A., Cornforth A.N., Kuznetsov Y., McPherson A., Martini F., Jadus M.R. Fascin-1 knock-down of human glioma cells reduces their microvilli/filopodia while improving their susceptibility to lymphocyte-mediated cytotoxicity. Am. J. Transl. Res. 2015;7:271-284.
Not in our library
527.
Eke I., Storch K., Kästner I., Vehlow A., Faethe C., Mueller-Klieser W., Taucher-Scholz G., Temme A., Schackert G., Cordes N. Three-dimensional invasion of human glioblastoma cells remains unchanged by X-ray and carbon ion irradiation in vitro. Int. J. Radiat. Oncol. Biol. Phys. 2012;84.
Not in our library10.1016/j.ijrobp.2012.06.012
528.
Frankel P., Pellet-Many C., Lehtolainen P., D’Abaco G.M., Tickner M.L., Cheng L., Zachary I.C. Chondroitin sulphate-modified neuropilin 1 is expressed in human tumour cells and modulates 3D invasion in the U87MG human glioblastoma cell line through a p130Cas-mediated pathway. EMBO Rep. 2008;9:983-989.
Not in our library10.1038/embor.2008.151
529.
Oppel F., Müller N., Schackert G., Hendruschk S., Martin D., Geiger K.D., Temme A. SOX2-RNAi attenuates S-phase entry and induces RhoA-dependent switch to protease-independent amoeboid migration in human glioma cells. Mol. Cancer. 2011;10:1-16.
Not in our library10.1186/1476-4598-10-137
530.
Weeks A., Okolowsky N., Golbourn B., Ivanchuk S., Smith C., Rutka J.T. ECT2 and RASAL2 mediate mesenchymal-amoeboid transition in human astrocytoma cells. Am. J. Pathol. 2012;181:662-674.
Not in our library10.1016/j.ajpath.2012.04.011
531.
Quick Q., Paul M., Skalli O. Roles and potential clinical applications of intermediate filament proteins in brain tumors. Semin. Pediatr. Neurol. 2015;22:40-48.
Not in our library10.1016/j.spen.2014.12.005
532.
Ducray F., Mokhtari K., Crinire E., Idbaih A., Marie Y., Dehais C., Paris S., Carpentier C., Dieme M.J., Adam C., et al. Diagnostic and prognostic value of alpha internexin expression in a series of 409 gliomas. Eur. J. Cancer. 2011;47:802-808.
Not in our library10.1016/j.ejca.2010.11.031
533.
Skalli O., Wilhelmsson U., Örndahl C., Fekete B., Malmgren K., Rydenhag B., Pekny M. Astrocytoma grade IV (glioblastoma multiforme) displays 3 subtypes with unique expression profiles of intermediate filament proteins. Hum. Pathol. 2013;44:2081-2088.
Not in our library10.1016/j.humpath.2013.03.013
534.
Paetau A. Glial fibrillary acidic protein, vimentin and fibronectin in primary cultures of human glioma and fetal brain. Acta Neuropathol. 1988;75:448-455.
Not in our library10.1007/bf00687131
535.
Yung W.A., Luna M., Borit A. Vimentin and Glial Fibrillary Acidic Protein in Human Brain Tumors. J. Neuro-Oncol. 1985;38:35-38.
Not in our library10.1007/bf00165169
536.
van Bodegraven E.J., van Asperen J.V., Robe P.A.J., Hol E.M. Importance of GFAP isoform-specific analyses in astrocytoma. Glia. 2019;1-17.
Not in our library10.1002/glia.23594
537.
Lin L., Wang G., Ming J., Meng X., Han B., Sun B., Cai J., Jiang C. Analysis of expression and prognostic significance of vimentin and the response to temozolomide in glioma patients. Tumor Biol. 2016;37:15333-15339.
Not in our library10.1007/s13277-016-5462-7
538.
Zhao J., Zhang L., Dong X., Liu L., Huo L., Chen H. High Expression of Vimentin is Associated with Progression and a Poor Outcome in Glioblastoma. Appl. Immunohistochem. Mol. Morphol. 2018;26:337-344.
Not in our library10.1097/pai.0000000000000420
539.
Mahesparan R., Read T.-A., Lund-Johansen M., Skaftnesmo K.O., Bjerkvig R., Engebraaten O. Expression of extracellular matrix components in a highly infiltrative in vivo glioma model. Acta Neuropathol. 2003;105:49-57.
Not in our library10.1007/s00401-002-0610-0
540.
Lv D., Lu L., Hu Z., Fei Z., Liu M., Wei L., Xu J. Nestin Expression Is Associated with Poor Clinicopathological Features and Prognosis in Glioma Patients: An Association Study and Meta-analysis. Mol. Neurobiol. 2017;54:727-735.
Not in our library10.1007/s12035-016-9689-5
541.
Baumgarten P., Quick-Weller J., Gessler F., Wagner M., Tichy J., Forster M.T., Foerch C., Seifert V., Mittelbronn M., Senft C. Pre- and early postoperative GFAP serum levels in glioma and brain metastases. J. Neurooncol. 2018;139:541-546.
Not in our library10.1007/s11060-018-2898-1
542.
Elobeid A., Bongcam-Rudloff E., Westermark B., Nistér M. Effects of inducible glial fibrillary acidic protein on glioma cell motility and proliferation. J. Neurosci. Res. 2000;60:245-256.
Not in our library10.1002/(sici)1097-4547(20000415)60:2<245::aid-jnr14>3.0.co;2-1
543.
Toda M., Miura M., Asou H., Toya S., Uyemura K. Cell growth suppression of astrocytoma C6 cells by glial fibrillary acidic protein cDNA transfection. J. Neurochem. 1994;63:1975-1978.
Not in our library10.1046/j.1471-4159.1994.63051975.x
544.
Kouam P.N., Rezniczek G.A., Kochanneck A., Priesch-Grzeszkowiak B., Hero T., Adamietz I.A., Bühler H. Robo1 and vimentin regulate radiation-induced motility of human glioblastoma cells. PLoS ONE. 2018;13:1-18.
Not in our library10.1371/journal.pone.0198508
545.
Ishiwata T., Teduka K., Yamamoto T., Kawahara K., Matsuda Y., Naito Z. Neuroepithelial stem cell marker nestin regulates the migration, invasion and growth of human gliomas. Oncol. Rep. 2011;26:91-99.
Not in our library10.3892/or.2011.1267
546.
Kitai R., Horita R., Sato K., Yoshida K., Arishima H., Higashino Y., Hashimoto N., Takeuchi H., Kubota T., Kikuta K.-I. Nestin expression in astrocytic tumors delineates tumor infiltration. Brain Tumor Pathol. 2010;27:17-21.
Not in our library10.1007/s10014-009-0261-0
547.
Jing R., Pizzolato G., Robson R.M., Gabbiani G., Skalli O. Intermediate filament protein synemin is present in human reactive and malignant astrocytes and associates with ruffled membranes in astrocytoma cells. Glia. 2005;50:107-120.
Not in our library10.1002/glia.20158
548.
Rickman D.S., Bobek M.P., Misek D.E., Kuick R., Blaivas M., Kurnit D.M., Taylor J., Hanash S.M. Distinctive molecular profiles of high-grade and low-grade gliomas based on oligonucleotide microarray analysis. Cancer Res. 2001;61:6885-6891.
Not in our library
549.
Katsetos C.D., Legido A., Perentes E., Mörk S.J. Class III β-Tubulin Isotype: A Key Cytoskeletal Protein at the Crossroads of Developmental Neurobiology and Tumor Neuropathology. J. Child Neurol. 2003;18:851-866.
Not in our library10.1177/088307380301801205
550.
Katsetos C.D., Reddy G., Dráberová E., Šmejkalová B., Del Valle L., Ashraf Q., Tadevosyan A., Yelin K., Maraziotis T., Mishra O.P., et al. Altered Cellular Distribution and Subcellular Sorting of γ-Tubulin in Diffuse Astrocytic Gliomas and Human Glioblastoma Cell Lines. J. Neuropathol. Exp. Neurol. 2006;65:465-477.
Not in our library10.1097/01.jnen.0000229235.20995.6e
551.
Katsetos C.D., Reginato M.J., Baas P.W., D’Agostino L., Legido A., Tuszyński J.A., Dráberová E., Dráber P. Emerging microtubule targets in glioma therapy. Semin. Pediatr. Neurol. 2015;22:49-72.
Not in our library10.1016/j.spen.2015.03.009
552.
Orr G.A., Verdier-Pinard P., McDaid H., Horwitz S.B. Mechanisms of Taxol resistance related to microtubules. Oncogene. 2003;22:7280-7295.
Not in our library10.1038/sj.onc.1206934
553.
Kavallaris M., Kuo D.Y., Burkhart C.A., Regl D.L., Norris M.D., Haber M., Horwitz S.B. Taxol-resistant epithelial ovarian tumors are associated with altered expression of specific beta-tubulin isotypes. J. Clin. Investig. 1997;100:1282-1293.
Not in our library10.1172/jci119642
554.
Karmakar S., Banik N.L., Ray S.K. Combination of all-trans retinoic acid and paclitaxel-induced differentiation and apoptosis in human glioblastoma U87MG xenografts in nude mice. Cancer. 2008;112:596-607.
Not in our library10.1002/cncr.23223
555.
Chao C.C., Kan D., Lo T.H., Lu K.S., Chien C.-L. Induction of neural differentiation in rat C6 glioma cells with taxol. Brain Behav. 2015;5:1-10.
Not in our library10.1002/brb3.414
556.
Godinho S.A., Pellman D. Causes and consequences of centrosome abnormalities in cancer. Philos. Trans. R. Soc. Lond. B Biol. Sci. 2014;369.
Not in our library10.1098/rstb.2013.0467
557.
Dráberová E., Vinopal S., Morfini G., Liu P.S., Sládková V., Sulimenko T., Burns M.R., Solowska J., Kulandaivel K., De Chadarévian J.P., et al. 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. 2011;70:811-826.
Not in our library10.1097/nen.0b013e31822c256d
558.
Song Y., Mu L., Han X., Liu X., Fu S. SiRNA targeting stathmin inhibits invasion and enhances chemotherapy sensitivity of stem cells derived from Glioma cell lines. Acta Biochim. Biophys. Sin. (Shanghai). 2014;46:1034-1040.
Not in our library10.1093/abbs/gmu099
559.
Masui K., Mawatari S., Suzuki S.O., Iwaki T. Evaluation of sensitivity and specificity of doublecortin immunostatining for the detection of infiltrating glioma cells. Brain Tumor Pathol. 2008;25:1-7.
Not in our library10.1007/s10014-007-0225-1
560.
Daou M.-C., Smith T.W., Litofsky N.S., Hsieh C.C., Ross A.H. Doublecortin is preferentially expressed in invasive human brain tumors. Acta Neuropathol. 2005;110:472-480.
Not in our library10.1007/s00401-005-1070-0
561.
Santra M., Santra S., Roberts C., Zhang R.L., Chopp M. Doublecortin induces mitotic microtubule catastrophe and inhibits glioma cell invasion. J. Neurochem. 2009;108:231-245.
Not in our library10.1111/j.1471-4159.2008.05758.x
562.
Santra M., Zhang X., Santra S., Jiang F., Chopp M. Ectopic Doublecortin Gene Expression Suppresses the Malignant Phenotype in Glioblastoma Cells. Cancer Res. 2006;66:11726-11735.
Not in our library10.1158/0008-5472.can-06-1978
563.
Pagano A., Honoré S., Mohan R., Berges R., Akhmanova A., Braguer D. Epothilone B inhibits migration of glioblastoma cells by inducing microtubule catastrophes and affecting EB1 accumulation at microtubule plus ends. Biochem. Pharmacol. 2012;84:432-443.
Not in our library10.1016/j.bcp.2012.05.010
564.
Näthke I.S. THE ADENOMATOUS POLYPOSIS COLI PROTEIN: The Achilles Heel of the Gut Epithelium. Annu. Rev. Cell Dev. Biol. 2004;20:337-366.
Not in our library10.1146/annurev.cellbio.20.012103.094541
565.
Segditsas S., Tomlinson I. Colorectal cancer and genetic alterations in the Wnt pathway. Oncogene. 2006;25:7531-7537.
Not in our library10.1038/sj.onc.1210059
566.
Suzuki S.O., McKenney R.J., Mawatari S.Y., Mizuguchi M., Mikami A., Iwaki T., Goldman J.E., Canoll P., Vallee R.B. Expression patterns of LIS1, dynein and their interaction partners dynactin, NudE, NudEL and NudC in human gliomas suggest roles in invasion and proliferation. Acta Neuropathol. 2007;113:591-599.
Not in our library10.1007/s00401-006-0180-7
567.
Zhang J., Guo W.-H., Wang Y.-L. Microtubules stabilize cell polarity by localizing rear signals. Proc. Natl. Acad. Sci. USA. 2014;111:16383-16388.
Not in our library10.1073/pnas.1410533111
568.
Bie L., Zhao G., Wang Y.P., Zhang B. Kinesin family member 2C (KIF2C/MCAK) is a novel marker for prognosis in human gliomas. Clin. Neurol. Neurosurg. 2012;114:356-360.
Not in our library10.1016/j.clineuro.2011.11.005
569.
Wang Q., Wang L., Li D., Deng J., Zhao Z., He S., Zhang Y., Tu Y. Kinesin family member 14 is a candidate prognostic marker for outcome of glioma patients. Cancer Epidemiol. 2013;37:79-84.
Not in our library10.1016/j.canep.2012.08.011
570.
Venere M., Horbinski C., Crish J.F., Jin X., Vasanji A., Major J., Burrows A.C., Chang C., Prokop J., Wu Q., et al. The mitotic kinesin KIF11 is a driver of invasion, proliferation, and self-renewal in glioblastoma. Sci. Transl. Med. 2015;7:304ra143.
Not in our library10.1126/scitranslmed.aac6762
571.
Afghani N., Mehta T., Wang J., Tang N., Skalli O., Quick Q.A. Microtubule actin cross-linking factor 1, a novel target in glioblastoma. Int. J. Oncol. 2017;50:310-316.
Not in our library10.3892/ijo.2016.3798
572.
Arden J.D., Lavik K.I., Rubinic K.A., Chiaia N., Khuder S.A., Howard M.J., Nestor-Kalinoski A.L., Alberts A.S., Eisenmann K.M. Small-molecule agonists of mammalian Diaphanous-related (mDia) formins reveal an effective glioblastoma anti-invasion strategy. Mol. Biol. Cell. 2015;26:3704-3718.
Not in our library10.1091/mbc.e14-11-1502
573.
Hiratsuka M., Inoue T., Toda T., Kimura N., Shirayoshi Y., Kamitani H., Watanabe T., Ohama E., Tahimic C.G.T., Kurimasa A., et al. Proteomics-based identification of differentially expressed genes in human gliomas: Down-regulation of SIRT2 gene. Biochem. Biophys. Res. Commun. 2003;309:558-566.
Not in our library10.1016/j.bbrc.2003.08.029
574.
Gene Expression Profile of Glioblastoma Multiforme Invasive Phenotype Points to New Therapeutic Targets · 2005
Not checked
575.
Geiger K.D., Stoldt P., Schlote W., Derouiche A. Ezrin Immunoreactivity Is Associated with Increasing Malignancy of Astrocytic Tumors but Is Absent in Oligodendrogliomas. Am. J. Pathol. 2000;157:1785-1793.
Not in our library10.1016/s0002-9440(10)64816-x
576.
Tynninen O., Carpen O., Jaaskelainen J., Paavonen T., Paetau A. Ezrin expression in tissue microarray of primary and recurrent gliomas. Neuropathol. Appl. Neurobiol. 2004;30:472-477.
Not in our library10.1111/j.1365-2990.2004.00562.x
577.
Belot N., Rorive S., Doyen I., Lefranc F., Bruyneel E., Dedecker R., Micik S., Brotchi J., Decaestecker C., Salmon I., et al. Molecular characterization of cell substratum attachments in human glial tumors relates to prognostic features. Glia. 2001;36:375-390.
Not in our library10.1002/glia.1124
578.
Adams J.C. Roles of fascin in cell adhesion and motility. Curr. Opin. Cell Biol. 2004;16:590-596.
Not in our library10.1016/j.ceb.2004.07.009
579.
Peraud A., Mondal S., Hawkins C., Mastronardi M., Bailey K., Rutka J.T. Expression of fascin, an actin-bundling protein, in astrocytomas of varying grades. Brain Tumor Pathol. 2003;20:53-58.
Not in our library10.1007/bf02483447
580.
Ng H.K., Ko H.C.W., Tse C.C.H. Immunohistochemical and Ultrastructural Studies of Oligodendrogliomas Revealed Features of Neuronal Differentiation. Int. J. Surg. Pathol. 1994;2:47-55.
Not in our library10.1177/106689699400200109
581.
Wharton S.B., Chan K.K., Whittle I.R. Microtubule-associated protein 2 (MAP-2) is expressed in low and high grade diffuse astrocytomas. J. Clin. Neurosci. 2002;9:165-169.
Not in our library10.1054/jocn.2001.1055
582.
Zhang Y., Ni S., Huang B., Wang L., Zhang X., Li X., Wang H., Liu S., Hao A., Li X. Overexpression of SCLIP promotes growth and motility in glioblastoma cells. Cancer Biol. Ther. 2015;16:97-105.
Not in our library10.4161/15384047.2014.987037
583.
Jin X., Jin X., Jung J.E., Beck S., Kim H. Cell surface Nestin is a biomarker for glioma stem cells. Biochem. Biophys. Res. Commun. 2013;433:496-501.
Not in our library10.1016/j.bbrc.2013.03.021
584.
Strojnik T., Røsland G.V., Sakariassen P.O., Kavalar R., Lah T. Neural stem cell markers, nestin and musashi proteins, in the progression of human glioma: Correlation of nestin with prognosis of patient survival. Surg. Neurol. 2007;68:133-143.
Not in our library10.1016/j.surneu.2006.10.050
585.
Dehghani F., Schachenmayr W., Laun A., Korf H.W. Prognostic implication of histopathological, immunohistochemical and clinical features of oligodendrogliomas: A study of 89 cases. Acta Neuropathol. 1998;95:493-504.
Not in our library10.1007/s004010050830
586.
Forget M.-A., Desrosiers R.R., Del M., Moumdjian R., Shedid D., Berthelet F., Béliveau R. The expression of rho proteins decreases with human brain tumor progression: Potential tumor markers. Clin. Exp. Metastasis. 2002;19:9-15.
Not in our library10.1023/a:1013884426692
587.
Kwiatkowska A., Didier S., Fortin S., Chuang Y., White T., Berens M.E., Rushing E., Eschbacher J., Tran N.L., Chan A., et al. The small GTPase RhoG mediates glioblastoma cell invasion. Mol. Cancer. 2012;11:65.
Not in our library10.1186/1476-4598-11-65
588.
Rutka J.T., Muller M., Hubbard S.L., Forsdike J., Dirks P.B., Jung S., Tsugu A., Ivanchuk S., Costello P., Mondal S., et al. Astrocytoma Adhesion to Extracellular Matrix: Functional Significance of Integrin and Focal Adhesion Kinase Expression. J. Neuropathol. Exp. Neurol. 1999;58:198-209.
Not in our library10.1097/00005072-199902000-00009
589.
Natarajan M., Hecker T.P., Gladson C.L. FAK signaling in anaplastic astrocytoma and glioblastoma tumors. Cancer J. 2003;9:126-133.
Not in our library10.1097/00130404-200303000-00008
590.
Mariani L., Beaudry C., McDonough W.S., Hoelzinger D.B., Demuth T., Ross K.R., Berens T., Coons S.W., Watts G., Trent J.M., et al. Glioma cell motility is associated with reduced transcription of proapoptotic and proliferation genes: A cDNA microarray analysis. J. Neurooncol. 2001;53:161-176.
Not in our library10.1023/a:1012253317934
591.
Giese A., Loo A., Norman S.A., Treasurywala S., Berens M.E. Contrasting migratory response of astrocytoma cells to tenascin mediated by different integrins. J. Cell Sci. 1996;109:2161-2168.
Not in our library10.1242/jcs.109.8.2161
592.
Plopper G.E., McNamee H.P., Dike L.E., Bojanowski K., Ingber D.E. Convergence of integrin and growth factor receptor signaling pathways within the focal adhesion complex. Mol. Biol. Cell. 1995;6:1349-1365.
Not in our library10.1091/mbc.6.10.1349
593.
Wenk M.B., Midwood K.S., Schwarzbauer J.E. Tenascin-C suppresses Rho activation. J. Cell Biol. 2000;150:913-920.
Not in our library10.1083/jcb.150.4.913
594.
Hirata E., Arakawa Y., Shirahata M., Yamaguchi M., Kishi Y., Okada T., Takahashi J.A., Matsuda M., Hashimoto N. Endogenous tenascin-C enhances glioblastoma invasion with reactive change of surrounding brain tissue. Cancer Sci. 2009;100:1451-1459.
Not in our library10.1111/j.1349-7006.2009.01189.x
595.
Platten M., Wick W., Wild-Bode C., Aulwurm S., Dichgans J., Weller M. Transforming Growth Factors β1 (TGF-β1) and TGF-β2 Promote Glioma Cell Migration via Up-Regulation of αVβ3 Integrin Expression. Biochem. Biophys. Res. Commun. 2000;268:607-611.
Not in our library10.1006/bbrc.2000.2176
596.
Tonn J.C., Wunderlich S., Kerkau S., Klein C.E., Roosen K. Invasive behaviour of human gliomas is mediated by interindividually different integrin patterns. Anticancer Res. 1998;18:2599-2605.
Not in our library
597.
Rooprai H.K., Vanmeter T., Panou C., Schnüll S., Trillo-Pazos G., Davies D., Pilkington G.J. The role of integrin receptors in aspects of glioma invasion in vitro. Int. J. Dev. Neurosci. 1999;17:613-623.
Not in our library10.1016/s0736-5748(99)00051-9
598.
Gritsenko P.G., Friedl P. Adaptive adhesion systems mediate glioma cell invasion in complex environments. J. Cell Sci. 2018;131:jcs216382.
Not in our library10.1242/jcs.216382
599.
Serres E., Debarbieux F., Stanchi F., Maggiorella L., Grall D., Turchi L., Burel-Vandenbos F., Figarella-Branger D., Virolle T., Rougon G., et al. Fibronectin expression in glioblastomas promotes cell cohesion, collective invasion of basement membrane in vitro and orthotopic tumor growth in mice. Oncogene. 2014;33:3451-3462.
Not in our library10.1038/onc.2013.305
600.
Demuth T., Berens M.E. Molecular mechanisms of glioma cell migration and invasion. J. Neurooncol. 2004;70:217-228.
Not in our library10.1007/s11060-004-2751-6
601.
Giese A., Kluwe L., Laube B., Meissner H., Berens M.E., Westphal M. Migration of human glioma cells on myelin. Neurosurgery. 1996;38:755-764.
Not in our library10.1227/00006123-199604000-00026
602.
Lash L.L., Wallar B.J., Turner J.D., Vroegop S.M., Kilkuskie R.E., Kitchen-Goosen S.M., Xu H.E., Alberts A.S. Small-Molecule Intramimics of Formin Autoinhibition: A New Strategy to Target the Cytoskeletal Remodeling Machinery in Cancer Cells. Cancer Res. 2013;73:6793-6803.
Not in our library10.1158/0008-5472.can-13-1593
603.
Wiranowska M., Ladd S., Smith S.R., Gottschall P.E. CD44 adhesion molecule and neuro-glial proteoglycan NG2 as invasive markers of glioma. Brain Cell Biol. 2006;35:159-172.
Not in our library10.1007/s11068-007-9009-0
604.
Akiyama Y., Jung S., Salhia B., Lee S., Hubbard S., Taylor M., Mainprize T., Akaishi K., van Furth W., Rutka J.T. Hyaluronate Receptors Mediating Glioma Cell Migration and Proliferation. J. Neurooncol. 2001;53:115-127.
Not in our library10.1023/a:1012297132047
605.
Toole B.P. Hyaluronan: From extracellular glue to pericellular cue. Nat. Rev. Cancer. 2004;4:528-539.
Not in our library10.1038/nrc1391
606.
Merzak A., Koocheckpour S., Pilkington G.J. CD44 mediates human glioma cell adhesion and invasion in vitro. Cancer Res. 1994;54:3988-3992.
Not in our library
607.
Park J.B., Kwak H.-J., Lee S.-H. Role of hyaluronan in glioma invasion. Cell Adh. Migr. 2008;2:202-207.
Not in our library10.4161/cam.2.3.6320
608.
Karousou E., Misra S., Ghatak S., Dobra K., Götte M., Vigetti D., Passi A., Karamanos N.K., Skandalis S.S. Roles and targeting of the HAS/hyaluronan/CD44 molecular system in cancer. Matrix Biol. 2017;59:3-22.
Not in our library10.1016/j.matbio.2016.10.001
609.
Bourguignon L.Y.W. Hyaluronan-mediated CD44 activation of RhoGTPase signaling and cytoskeleton function promotes tumor progression. Semin. Cancer Biol. 2008;18:251-259.
Not in our library10.1016/j.semcancer.2008.03.007
610.
Park D., Kim Y., Kim H., Kim K., Lee Y.-S., Choe J., Hahn J.-H., Lee H., Jeon J., Choi C., et al. Hyaluronic acid promotes angiogenesis by inducing RHAMM-TGFβ receptor interaction via CD44-PKCδ. Mol. Cells. 2012;33:563-574.
Not in our library10.1007/s10059-012-2294-1
611.
Murai T., Miyazaki Y., Nishinakamura H., Sugahara K.N., Miyauchi T., Sako Y., Yanagida T., Miyasaka M. Engagement of CD44 Promotes Rac Activation and CD44 Cleavage during Tumor Cell Migration. J. Biol. Chem. 2004;279:4541-4550.
Not in our library10.1074/jbc.m307356200
612.
Klank R.L., Decker Grunke S.A., Bangasser B.L., Forster C.L., Price M.A., Odde T.J., SantaCruz K.S., Rosenfeld S.S., Canoll P., Turley E.A., et al. Biphasic Dependence of Glioma Survival and Cell Migration on CD44 Expression Level. Cell Rep. 2017;18:23-31.
Not in our library10.1016/j.celrep.2016.12.024

Cited by papers we hold 1

Subdiffraction-resolution fluorescence imaging of immunological synapse formation between NK cells and A. fumigatus by expansion microscopy · 2021
Not checked

Reads

Read by people on this page

None yet.

Checked or read by AI through naulon

None yet.

A person counts once a day per paper. We keep no cookie and no address: a reader is a hash that changes every day. Any month under five reads shows no number.

Are you an author?

Is this your paper?

Claim it to add your note and be paid when agents cite it. Free.