Using fly genetics to dissect the cytoskeletal machinery of neurons during axonal growth and maintenance

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Using fly genetics to dissect the cytoskeletalmachinery of neurons during axonal growth andmaintenance

Andreas Prokop*, Robin Beaven, Yue Qu and Natalia Sanchez-Soriano*Faculty of Life Sciences, The University of Manchester, Michael Smith Building, Oxford Road, Manchester M13 9PT, UK

*Authors for correspondence ([email protected]; [email protected])

Journal of Cell Science 126, 2331–2341� 2013. Published by The Company of Biologists Ltddoi: 10.1242/jcs.126912

SummaryThe extension of long slender axons is a key process of neuronal circuit formation, both during brain development and regeneration. Forthis, growth cones at the tips of axons are guided towards their correct target cells by signals. Growth cone behaviour downstream of

these signals is implemented by their actin and microtubule cytoskeleton. In the first part of this Commentary, we discuss thefundamental roles of the cytoskeleton during axon growth. We present the various classes of actin- and microtubule-binding proteins thatregulate the cytoskeleton, and highlight the important gaps in our understanding of how these proteins functionally integrate into the

complex machinery that implements growth cone behaviour. Deciphering such machinery requires multidisciplinary approaches,including genetics and the use of simple model organisms. In the second part of this Commentary, we discuss how the application ofcombinatorial genetics in the versatile genetic model organism Drosophila melanogaster has started to contribute to the understanding

of actin and microtubule regulation during axon growth. Using the example of dystonin-linked neuron degeneration, we explain howknowledge acquired by studying axonal growth in flies can also deliver new understanding in other aspects of neuron biology, such asaxon maintenance in higher animals and humans.

Key words: Growth cone, Cytoskeleton, Axon, Drosophila, Actin, Microtubule, Brain disorders

IntroductionThe cytoskeleton consists of actin, intermediate filaments (IFs) and

microtubules (MTs), and has fundamental roles in virtually every

function of cells, including cell division, motility, adhesion, signalling,

endocytic trafficking and transport, as well as in the regulation of cell

and organelle shapes and their distributions (Akhmanova and Stearns,

2013). Naturally, this also applies to the nervous system where the

cytoskeleton has essential roles during the development, function,

regeneration and degeneration of neurons. Of the more than 200 entries

on the OMIM (Online Mendelian Inheritance in Man) database for

genes encoding cytoskeleton-associated proteins, 44% have indexed

links to human disorders, and 54% of these are linked to nervous

system disorders, including neurodevelopmental disorders (e.g.

lissencephalies, mental retardations), functional disorders (e.g.

deafness) and a wide range of degenerative diseases (see

supplementary material Table S1). Therefore, research of the

cytoskeleton provides unique opportunities to unravel fundamental

mechanisms of the nervous system, both in health and disease.

It is currently little understood how cytoskeleton-associated

proteins contribute to cellular mechanisms of neurons. For

example, the systemic importance of certain MT-binding

proteins, such as motor proteins and the structural protein tau,

has long been recognised from their involvement in a broad

spectrum of neurodegenerative diseases, and we know their

principal molecular functions. However, we have yet to explain

conclusively how they function within normal or diseased

neurons (Hirokawa et al., 2010; Morris et al., 2011). Here, we

argue that this is best performed starting with processes for which

there is already an extensive body of knowledge about the

cytoskeleton, in particular the growth of axons.

Axons are slender neuronal extensions, which can be several

metres long, and are often arranged into nerves or nerve tracts

that provide essential ‘information highways’ in the nervous

system. The proper function of nervous systems requires axons to

grow and wire up correctly during development or regeneration,

and these connections need to be maintained for decades in the

ageing body. Here, we review the current concepts for the

cellular roles that actin and MTs have in axons and discuss

important gaps in our understanding of how the cytoskeleton is

regulated to this end. We argue that genetics provides important

means to advance this understanding, and explain how studies

using the genetic model organism Drosophila melanogaster can

make important contributions. We review recent advances with

regard to actin and MT regulation during axonal growth in the fly

and provide an example of how mechanistic understanding in one

context can be used to identify mechanisms underpinning other

processes, such as axon maintenance and neurodegeneration.

The organisation of the cytoskeleton in axonsMature axons are the longest cellular entities that are produced by

animals. They propagate electrical messages, which travel from

the neuronal cell bodies towards the synaptic connections with

their distant target cells, often several metres away. To maintain

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this unusual architecture, shafts of axons contain prominent

bundles of parallel MTs, which provide structural support as well

as the tracks for the vital long-distance cargo transport

(Twelvetrees et al., 2012). Axonal MTs are discontinuous and

are usually not attached to the centrosome (the main microtubule

organizing centre of most cells); their plus ends mostly, but not

entirely, point distally (Fig. 1A) (Ahmad et al., 1999; Baas and

Lin, 2011; Basto et al., 2006; Stiess et al., 2010). The cortex of

axon shafts is lined by short stabilised actin filaments (Fig. 1A),

which are organised into repetitive rings by the scaffolding

protein spectrin and linked to the axonal cell membrane through

ankyrins (Xu et al., 2013). Axons also contain a number of

different IF proteins that can regulate the specific diameters of

different axon classes (Perrot and Eyer, 2009).

Axons are usually maintained for the lifetime of an organism,

i.e. for decades in humans. It is therefore not surprising that axonal

decay is a characteristic feature in the ageing brain (Marner et al.,

2003). Sites of severe disorganisation of axonal MT bundles

(referred to as diverticula) are considered an important trigger of

such axon loss (Adalbert et al., 2009; Fiala et al., 2007). Axonal

MT bundles are believed to be stabilised through structural MT-

binding proteins (MTBPs), such as tau or MAP1B (Chilton and

Gordon-Weeks, 2007). At the same time, there seems to be a

steady and dynamic MT turnover, as suggested by continued MT

polymerisation events in mature axons (Kollins et al., 2009). Such

dynamics might allow for structural plasticity, such as shortening

or extension of the axon shaft (Kuppers-Munther et al., 2004;

Lamoureux et al., 2010; Rajagopalan et al., 2010), pruning and

regrowth of axons or formation of new side branches (Letourneau,

2009), which are required for rewiring processes during learning

and memory formation or the regeneration of damaged axons

(Bradke et al., 2012; Saxena and Caroni, 2007).

Cellular roles of the cytoskeleton during axonaldevelopmentDuring development, axons are initiated as small neuritic

protrusions on the surface of neuronal cell bodies. For this,

MTs in the neuronal cell bodies become stabilised and push

beyond the cell cortex (Brandt, 1998). This is assisted by

dynamic rearrangements of the cortical actin cytoskeleton to

remove barrier function (Flynn et al., 2012; Neukirchen and

Bradke, 2011; Saengsawang et al., 2012) and the formation of

filopodia, which provide membrane protrusions into which MTs

can extend (Dent et al., 2007; Goncalves-Pimentel et al., 2011).

In typical vertebrate neurons, several neurites are formed

simultaneously on the cell body, but only one of these is

selected as the axon. This selection process (referred to as neuron

polarisation) involves signalling cascades leading to the targeted

Filopodia

Engorgingfilopodium

Engorging filopodium

Cortical actinCortical actin rings

F-actinlattice

F-actinbundle

F-actinarc

LamellipodiumSoma

Axon

CentrosomeMT polymerisation

F-actin backflowRepulsive cue

MT-destabi-lising drug

MT-stabilising drugElectric field MTF-actin

latticeF-actinbundle

StabilisedMT

Attractive cue

oror

or or

Growthcone

Nucleus

A

B

E F-act– F F-act– C Splayed MT– D F-act–

New axon segment

Key

Fig. 1. Principal organisation and roles of actin and

MTs in growing axons. (A) The figure shows the

principal distribution and organisation of MTs and F-

actin networks in the soma, axon and growth cone of a

neuron that is exposed to an attractive guidance cue.

The tip of the axonal MT bundle forms the central zone

of the growth cone from where splayed MTs emanate

into the actin-rich peripheral zone (Dent et al., 2011).

(B) Directed axon extension (growth cone turning).

Splayed MTs become preferentially stabilised in a

certain direction through guidance cues [or

experimentally through unilateral exposure to

MT-(de-)stabilising drugs or the application of an

electrical field]. Additional MTs follow in that

direction, leading to engorgement of membrane

structures in that area and, eventually, the establishment

of a new axon segment (Buck and Zheng, 2002;

Lowery and Van Vactor, 2009; McCaig et al., 2002;

Sabry et al., 1991; Tanaka and Kirschner, 1995;

Tanaka and Kirschner, 1991). (C) Growth cone turning

requires splayed MTs: low doses of MT-stabilising

drugs suppress the occurrence of splayed, side-stepping

MTs (splayed MT2). Growth cones that are treated this

way fail to show normal turning behaviour at repulsive

borders and stall instead (Challacombe et al., 1997;

Williamson et al., 1996). (D,E) Growth cone turning

requires F-actin. When F-actin is destabilised through

drugs (F-act2), MTs tend to be bundled and growth

cones fail to turn at repulsive borders (Challacombe et

al., 1996); they also no longer respond to unidirectional

application of MT-stabilising drugs (Buck and Zheng,

2002). (F) Growth cone turning towards the cathode in

electrical fields (galvanotropism) does not require F-

actin (McCaig, 1989); this turning could be mediated

by MTBPs, which are attracted to the cathode through

positive net-charge of their MT-binding domains, thus

dragging the MTs along with them.

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localisation of cell polarity factors (Barnes and Polleux, 2009)and also depends on the specific stabilisation of MTs in the

selected neurite (Brandt, 1998; Chen et al., 2013).

Once established, axons grow along stereotypic paths to theirtarget cells, a process that is implemented by prominent hand-shaped growth cones at the axon tips. Growth cones are guided

towards their specific target areas and cells through precisespatio-temporal patterns of chemical signals that are arrangedalong their paths (Tessier-Lavigne and Goodman, 1996). Many

guidance cues and their receptors have been identified and shownto mediate repulsion or attraction (Araujo and Tear, 2003; Huberet al., 2003), thus controlling growth cone behaviours, such as

growth velocity, pausing, turning, retraction or collapse. Thesemorphogenetic movements downstream of guidance signallingare implemented by the prominent actin and MT cytoskeleton ofgrowth cones (Fan et al., 1993), which is arranged in a particular

spatial geometry.

The geometry of growth cones can be subdivided into an actin-rich peripheral zone and a MT-rich central zone (Fig. 1A), and

the area in which MTs and F-actin prominently overlap is oftenreferred to as the transition zone. The central zone of growthcones comprises the tips of the axonal MT bundles, and from here

single MTs splay out into the peripheral zone. The peripheralzone consists of actin-rich membrane protrusions, which includefinger-like filopodia (containing parallel F-actin bundles) andveil-like lamellipodia (containing F-actin lattices; Fig. 1A).

Filopodia and lamellipodia sense guidance cues by presentingtheir receptors on their surfaces. They undergo constant shapechanges, driven by continuous assembly of actin filaments at the

leading edges, the retrograde flow of these filaments, and theirdepolymerisation and recycling further back in the transitionzone (Box 1). This actin dynamics allow filopodia to reach out

into the growth cone environment and act as signalling andadhesion sensors (Chien et al., 1993; Davenport et al., 1993;Dwivedy et al., 2007; Mattila and Lappalainen, 2008). They

allow lamellipodia to act as dynamic sites of substrate adhesionand myosin-II-driven contraction and force generation, which areimportant in influencing MT behaviours during guided axonextension (Box 2).

In current models of growth cone guidance (Fig. 1B) (Dentet al., 2011; Lowery and Van Vactor, 2009), external signalstrigger the directional stabilisation of single MTs in the

peripheral zone. These stabilised MTs are then joined by theextension of the bulk of MTs from the central domain (a processreferred to as engorgement), thus giving rise to a new axon

segment. This elongation step is completed when lamellipodiaand filopodia relocate distally to the newly formed axon tip(Fig. 1B). In this model, MTs are the essential implementers ofaxon extension, whereas F-actin is required for the directionality

of this extension (through influencing MT behaviours; Box 2).Accordingly, blocking MT dynamics inhibits axon growth(Letourneau and Ressler, 1984; Sanchez-Soriano et al., 2010;

Tanaka et al., 1995), whereas the pharmacological or geneticinhibition of actin polymerisation, either in cultured neurons or in

vivo, does not usually suppress axonal growth per se, but

abrogates growth cone turning and pathfinding (Fig. 1D,E) (Buckand Zheng, 2002; Challacombe et al., 1996; Chien et al., 1993;Kaufmann et al., 1998; Marsh and Letourneau, 1984; Sanchez-

Soriano et al., 2010). This said, a directional movement of growthcones does not always require F-actin (Fig. 1F), as somesignalling mechanisms can target MTs directly. For example,

GSK3-mediated phosphorylation of the MTBPs APC1, MAP1B

and/or CLASP is known to directly regulate MT stability in

growth cones (Hur et al., 2011; Lucas et al., 1998; Purro et al.,

2008).

Box 1. ABPs and MTBPs as essential regulators ofcytoskeletal dynamics

MTs are stiff hollow tubes typically formed by 13 protofilaments

(filamentous polymers of a- and b-tubulin heterodimers) that grow

and shrink primarily at their plus ends. Actin filaments are more

flexible double-helical chains of actin monomers that maintain their

equilibrium mainly by growth at plus ends and shrinkage at minus

ends (Fletcher and Mullins, 2010; Hawkins et al., 2010; Stricker

et al., 2010). Both have in common that they are polymers of

repetitive building blocks (black helmet-like symbols in the figure)

that are assembled in a polar fashion. Their dynamics are

regulated in comparable ways through the following different

classes of actin-binding proteins (ABPs) and MT-binding proteins

(MTBPs) as illustrated in the box figure.

(1) New actin filaments or MTs are generated through nucleation

factors that catalyse the transition from mono- or oligomers to

polymers; (2) plus-end dynamics, such as (de-)polymerisation,

stabilisation, directionality and targeting is regulated by plus-end-

binding proteins; (3) nucleation or (de-)polymerisation processes

are further regulated by proteins that bind actin or tubulin

monomers or oligomers and determine their availability, for

example, SCAR or N-WASP activates Arp2/3 nucleation, APC1

cooperates with mDia1 in nucleation, stathmin sequesters tubulin,

profilin enhances actin polymerisation by ENAH (Bear and Gertler,

2009; Okada et al., 2010; Pollitt and Insall, 2009; Steinmetz,

2007); (4) proteins that bind along actin filaments or MTs can

stabilise them against depolymerisation, cross-link them into

bundles and/or networks, and/or link them to other cytoskeletal

components, organelles or the cortex; (5) minus-end-binding

proteins regulate stability versus (de-)polymerisation; (6,7) plus-

and minus-end-directed motor proteins mediate filament

contraction, the sliding along other cellular structures or

filaments, or the transport of cargo (C in the figure) along F-actin

or MTs; (8) different classes of proteins can sever or actively

depolymerise F-actin or MTs, for example, cofilin depolymerises or

severs pointed ends of F-actin, type 13 kinesins depolymerise MTs

from the plus end, and katanin or spastin sever MT shafts (Pak

et al., 2008; Conde and Caceres, 2009); (9) post-translational

modifications (PTM) influence F-actin or MT stability and the

interaction with certain ABPs and MTBPs (Fukushima et al., 2009;

Janke and Bulinski, 2011; Terman and Kashina, 2013).

91

2

6

7

85

4

3

33

C

CPTM

Nucleation

G-actin orα-tubulin–β-tubulin

Polymer/network regulation

Key

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The importance and challenge of understandingcytoskeletal machineryClearly, the cytoskeleton has essential roles during axonal

growth, but we still do not understand how it is regulated to

perform these functions. Different classes of actin-binding

proteins (ABPs) and MTBPs are known to regulate cytoskeletal

dynamics (Box 2); many of these are expressed in neurons

(Table 1), and we often have a reasonable understanding of their

molecular mechanisms, at least in vitro. However, we do not

understand how these individual regulators functionally integrate

into the common cytoskeletal machinery that ultimately

implements neuronal behaviours. This deficit also leaves us

with little means to explain the cellular aberrations caused

by mutations of genes encoding ABPs or MTBPs that have

been linked to brain disorders, including disorders of

neurodevelopment and axonal growth (supplementary material

Table S1) (Nugent et al., 2012; Tischfield et al., 2011).

The need to understand cytoskeletal regulation at the cellular

level in neurons and non-neuronal cells alike is a current problem(Fletcher and Mullins, 2010; Insall and Machesky, 2009). This isclearly reflected by the number of review articles that attempt to

propose models explaining the functional integration ofcytoskeletal regulators (e.g. Conde and Caceres, 2009; Dentet al., 2011; Gallo, 2013; Lowery and Van Vactor, 2009; Paket al., 2008). For example, there are mechanistic models aiming

to explain how different classes of ABPs functionally combineduring filopodia formation (Mattila and Lappalainen, 2008) andmodels based on end-binding (EB) proteins that propose how an

increasing number of MT-plus-end-associating proteins canregulate MT dynamics (Akhmanova and Steinmetz, 2008;Akhmanova and Steinmetz, 2010) (see also Fig. 2). Currently,

such models depend on knowledge gained from a broad rangeof different neuronal systems, and they therefore remainhypothetical and are difficult to test experimentally.

In this Commentary, we promote strategic considerations that

will help to overcome some of the current shortcomings inexplaining cytoskeletal machinery. We believe that strongeremphasis must be given to studying the various parts of

cytoskeletal machinery consistently in the same neuronsystems, so that their functional links and interfaces can bedirectly assessed. These neuronal systems should not only be

amenable to biochemical, biophysical and cell biologicalapproaches, but also to the use of combinatorial genetics,therefore allowing a systematic assessment of different genemanipulations or combinations of mutations in the same cells.

Such a strategy will help us to integrate the knowledge we haveabout single cytoskeletal regulators so that we can develophigher-order concepts about the governance of the cytoskeletal

machinery during axon growth or in the context of braindisorders.

Drosophila as a model to study cytoskeletal machineryduring axonal growthOne organism in which combinatorial genetics has been andcontinues to be most successfully applied to the study of complex

biological problems is the fruit fly Drosophila melanogaster

(Keller, 1996; Roote and Prokop, 2013). Work in Drosophila islow cost, impressively fast, and particularly amenable to geneticmanipulations and unbiased genetic screens (Sanchez-Soriano

et al., 2007). Accordingly, Drosophila research has been anincubator for new ideas and concepts in neurobiology, and workin the fly revealed many fundamental molecular and cellular

mechanisms that have turned out to be conserved in higheranimals (Bellen et al., 2010). Work in Drosophila has madeimportant contributions to the understanding of axon growth,

including the discovery and analysis of proteins involved inpathfinding, and the discovery of mechanisms underpinningpioneer guidance, nerve branching, target cell recognition and

axonal transport (Araujo and Tear, 2003; Astigarraga et al., 2010;Brochtrup and Hummel, 2011; Landgraf and Thor, 2006;Sanchez-Soriano and Prokop, 2005; Sanchez-Soriano et al.,2007; Zala et al., 2013).

To study the cytoskeletal machinery during axon growth, flyneurons offer a number of useful features. First, most of theDrosophila ABPs and MTBPs display a high level of sequence

conservation with their mammalian counterparts, and they areusually expressed in the nervous system (Table 1) (Sanchez-Soriano et al., 2007). Second, cytoskeletal regulators tend to

Box 2. How F-actin networks influence MTdynamics in growth cones

N F-actin networks generate membrane protrusions on all sides

of growth cones that can be invaded by MTs in any direction.

This facilitates lateral extension and stabilisation of MTs as an

important prerequisite for growth cone turning (see also

Fig. 1B).

N F-actin networks can influence MT behaviours mechanically

(Lee and Suter, 2008; Schaefer et al., 2002) (see also Fig. 1A).

Thus, MT extension is antagonised by F-actin backflow or

through the formation of transverse bundles (arcs) that form

‘road-blocks’. MT extension can be promoted and guided

through the formation of radial F-actin bundles as tracks for MT

elongation. MTs can be pushed laterally or medially through

actomyosin contractions. Finally, MT extension can be

channelled into certain directions through coordinated loss of

F-actin from local spots within lamellipodia.

N Actin networks form scaffolds that serve as a platform for ABPs.

Some of these ABPs influence MT behaviours through mediating

actin–MT linkage. This linkage can be direct through ABPs that

also contain MT-binding domains [e.g. MAP1B, spectraplakins,

coronin 7 (POD1 in Drosophila), cytoplasmic-linker-associated

proteins (CLASPs) or adenomatous polyposis coli (APC)

(Bouquet et al., 2007; Moseley et al., 2007; Rothenberg et al.,

2003; Sanchez-Soriano et al., 2009; Tsvetkov et al., 2007)]. Other

ABPs are involved in indirect crosstalk with MTs by interacting

with MTBPs. Reported examples are the ABP drebrin that is

linked to the MTBP EB3, the ABP PPP1R9A (better known as

spinophilin) that binds to DCX, IQGAP that interacts with CLIP-

170, or functional interactions between the actin-binding motor

protein myosin II and the MT-associated dynein motor protein

complex (Bielas et al., 2007; Geraldo et al., 2008; Myers et al.,

2006; Swiech et al., 2011).

N Some ABPs can tether actin networks to transmembrane

receptors when they are engaged with their extracellular

ligands. This has been demonstrated for different classes of

transmembrane receptors (Giannone et al., 2009; Moore et al.,

2009; Moore et al., 2010). Such linkage of F-actin to the

extracellular environment generates mechanical forces across

the membrane, which can trigger intracellular signalling events

(referred to as the ‘clutch’ mechanism) that then can influence

actin and MT dynamics (Suter and Forscher, 2000; Suter and

Miller, 2011).

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display minimal redundancy in the fly genome. For example,

eliminating the function of Ena/VASP requires a triple knockout

in mouse, whereas only a single gene needs to be removed in

Drosophila (Goncalves-Pimentel et al., 2011; Kwiatkowski et al.,

2007). Third, mutant alleles or other genetic tools are often

readily available (McQuilton et al., 2012), and they can usually

be combined in the same animal in a short time (Goncalves-

Pimentel et al., 2011; Koper et al., 2012). Finally,

experimentation is fast; axon extension can be studied in fly

embryos after only half a day of development, and in primary

neurons after only 6 hours in culture (Prokop et al., 2012;

Sanchez-Soriano et al., 2007).

To study cytoskeletal regulation during axonal growth, genetic

manipulations of cytoskeletal regulators can be combined with a

broad spectrum of morphological readouts for axon growth in

vivo (Sanchez-Soriano et al., 2007), including live imaging of

cytoskeletal dynamics (Mattie et al., 2010; Pawson et al., 2008).

This repertoire can be further expanded when making

complementary studies in Drosophila primary neurons, which

give access to quantitative measurements of filopodia and

lamellipodia, retrograde F-actin flow, the organisation,

stabilisation and polymerisation rates of MTs, as well as axonal

growth rates (Prokop et al., 2012; Sanchez-Soriano et al., 2010).

Such readouts are crucial to unravel how ABPs and MTBPs

regulate the cytoskeleton in cells. Notably, cytoskeletal dynamics

in fly neurons are similar to those of other animal neuron

systems, indicating that the underlying regulation is fundamental

and well preserved (Sanchez-Soriano et al., 2010). This said,

there will be limitations when using neurons of this little insect

because some aspects are different in vertebrate neurons;

for example, the axon length and life time, the absence

of neurofilaments and myelination, and their unipolar

organisation, including the spike-initiating zone, which is

located at the base of dendrites away from the soma and not

near the cell body as is the axon initial segment in vertebrates

(Grubb and Burrone, 2010; Sanchez-Soriano et al., 2005). In the

following sections, we will provide examples of work on axonal

growth in the fly that has contributed new understanding of

cytoskeletal machinery.

Use of the fly to unravel new ABP functionsduring axonal growthWithin a few years, work in Drosophila growth cones has

identified at least four new ABPs as actin regulators in neurons,including Shot, DAAM, Psidin and Canoe. Shot is the only

Drosophila member of the spectraplakin family of actin- and

MT-binding proteins with two close homologues in mammals,

MACF1 (also known as ACF7) and dystonin (DST, also known

as BPAG1) (Suozzi et al., 2012). Both Shot and ACF7 promote

filopodia formation (Sanchez-Soriano et al., 2009). In Shot, this

function is relevant for axon guidance at the CNS midline, and it

requires binding of the two EF-hand domains to the putative

translational regulator Extra bases (Exba, also known as eIF5C)

(Fig. 2A) (Lee et al., 2007; Sanchez-Soriano et al., 2009), thus

potentially contributing to local translation events in growth

cones (Van Horck and Holt, 2008). DAAM (DAAM1 in

mammals) is one of four Drosophila formins (Diaphanous,

DAAM, Fhos and CG32138) reported to be present in the

nervous system (Pawson et al., 2008; Prokop et al., 2011;

Sanchez-Soriano et al., 2007). DAAM has now been shown to be

the essential formin in growth cones of embryonic neurons,

where it promotes filopodia formation and axonal pathfinding in

Table 1. Examples of evolutionary conserved MTBPs and ABPs in human and fly

Cytoskeleton regulator class MTBPs ABPs

Nucleators c-tubulin ring complex (cTub23C and other components);doublecortin* (DCX-EMAP)

Arp2/3 complex (Arpc1, Arp3, etc.); formins(DAAM, Fhos, CG32138, dia)

Plus-end-binding proteins MAPFRE/EB* (eb1); MT-plus-end-tracking proteins (+TIPs);doublecortin* (DCX-EMAP); CKAP5/CHTOG/XMAP215*(msps)

ENAH/VASP* (ena); formins (DAAM, Fhos,CG32138, dia); capping proteins* (cpa, cpb);adducins* (hts)

Mono- or oligomer-binding proteins Stathmin/SCG10* (stai); DPYSL2/CRMP2* (CRMP); CKAP5/CHTOG/XMAP215* (msps); doublecortin* (DCX-EMAP);tub-specific chaperone E* (tbce); CLASP* (chb)

WASF/SCAR/WAVE* (SCAR); WASL/N-WASP*(WASp); APC* (Apc); b4-thymosin/TMSB4X*(cib)a; profilin* (chic); twinfilin* (twf)

Shaft stabilisers and cross-linkers MAPT* (tau); MAP2* (tau); MAP1B* (futsch); MACF1*;dystonin* (shot)

Fascin* (sn); a-actinin* (Actn); plastin/fimbrin*(Fim); filamin* (cher); tropomyosin* (Tm1);moesin*, ezrin*, radixin* (Moe); a/b-spectrin* (a/b-Spec); drebrin* (CG10083); coronin* (coro,pod1); MLLT4/Afadin-6* (cno); CTTN*(Cortactin); PPP1R9A* (spinophilin)b

Minus-end stabilisers c-tubulin ring complex (cTub23C and other components);CAMSAP1* (Patronin)

Tropomodulin* (tmod)b; Arp2/3 complex (Arpc1,Arp66B, etc.)

Plus-end motors Type 1 kinesins (Khc, Klc, Pat1); type 2 kinesins (Klp64D,Klp68D, Kif3C, Kap3); type 3 kinesins (unc-104, Khc-73)

Myosin XVA* (Myo10A); myosin VA* (didum);myosin II (zip, sqh)

Minus-end motors cytoplasmic dynein/dynactin (Dhc64C, ctp, sw, Gl, etc.); type13 kinesins/KIF2C/MCAK* (Klp10A, Klp59C, Klp59D)

Myosin VI* (jar)

Severers, de-polymerisers katanin* (Kat60); spastin* (spas); type 13 kinesins/KIF2C/MCAK* (Klp10A, Klp59C, Klp59D)

Cofilin* (tsr); gelsolin* (Gel)

Post-translational modifications(enzymes not listed)c

GTP hydrolysis; acetylation; (poly-) glycylation; (poly-)glutamylation; de-tyrosination; de-glutamylation (D2 tubulin)

ATP hydrolysis; potentially acetylation, arginylation,phosphorylation, etc.

Some proteins fulfil more than one function and are repeated in different positions; terms highlighted with an asterisk are listed in OMIM, other terms use otherdescriptors for protein complexes or classes. Terms in brackets are the orthologous Drosophila genes with known functions and/or prominent expression in thenervous system, as indicated in FlyBase (McQuilton et al., 2012).

aCib represents a triple-repeat of B4-thymosin.bNot declared as orthologues in FlyBase in spite of partial sequence homologies.cAs described by Janke and Bulinski, 2011; Terman and Kashina, 2013.

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the central nervous system (CNS) (Goncalves-Pimentel et al.,

2011; Matusek et al., 2008). Psidin (known as NAA25 in

mammals) is the auxiliary subunit of the NatB acetylation

complex required for protein acetylation (Stephan et al., 2012). In

addition, it has been shown that Psidin can act outside this

complex as an ABP that regulates actin dynamics both in neurons

and in non-neuronal cells of Drosophila (Kim et al., 2011;

Stephan et al., 2012). These studies show that Psidin antagonises

F-actin stabilisation mediated by Tropomyosin 1 in lamellipodia,

which is required for the pathfinding of olfactory neurons in the

fly brain. Canoe and its mammalian homologue MLLT4 (also

known as afadin, AFD6) are PDZ-domain-containing ABPs that

localise to adherens junctions and have been shown to interact

with profilin (Boettner et al., 2000; Lorger and Moelling, 2006;

Sawyer et al., 2009). In Canoe-deficient embryos, axonal

pathfinding at the CNS midline is impaired, and this aberration

correlates with two growth cone phenotypes: an increase in

filopodia numbers and a failure to localise Robo proteins

(receptors that are important regulators of midline pathfinding)

to filopodia (Slovakova et al., 2012). In this scenario, Canoe

EB1Exba /eIF5C

Tau

DA

AM

Arp

2/3

Enab

led

Prof

ilin

CpA

+BPs

isin

Tm1

Shot

Shot

MAP1BGRDC-tail

MtLSEFSR-rodplakinCH

EB1MTsExba/eIF5CF-actin

Axon extension

FilopodiaLamellipodia

Cortex

Other MTs

ApcDyneinType 13 kinesinsCLIP190

CLASP

XMAPDCX STMN

Ssp2

Actin bindingMT binding

EB1 binding (+TIP)Tubulin binding

Other binding

Actin regulatingOther regulatingprocessesSub-machinery

2 Actin dynamics

1 MT plus-end dynamics3 MTshaft

stability

MT growth

α-tubulin– F-actin

B

A

Key

β-tubulin

Interactions

Fig. 2. Roles of Shot in MT regulation at the interface with other classes of ABPs and MTBPs. (A) The functional domains of Shot (below the structure) and

their reported molecular interactions (above the structure). CH, calponin homology domains; plakin, plakin-like domain; SR-rod, spectrin repeat rod; EF, EF-hand

motifs; GRD, Gas2-related domain; MtLS, microtubule tip localisation sequence; C-tail, C-terminal tail. (B) A close-up of a growth cone showing a single MT and

three different aspects of the cytoskeletal machinery (emboxed and numbered), all of which are being studied in fly neurons. (1) Regulation of MT plus-end

dynamics; it includes end-binding proteins (here EB1) which directly bind MT plus ends and recruit MT-plus-end-tracking proteins (+TIPs) (blue arrows; see

Akhmanova and Steinmetz, 2008; Etienne-Manneville, 2010). Shot has to compete with other +TIPs for binding to a limited pool of EB1; when bound to EB1, it

links MT plus ends to F-actin (via its N-terminal CH domains) and guides polymerising MT plus ends in the direction of axon growth. The proper localisation of

Shot at MT plus ends also requires direct association with MTs through its GRD and C-terminal tail. CLASPs and XMAP bind tubulin and promote MT

polymerisation; CLASPs are +TIPs whereas XMAP215 (XMAP) binds MT plus-ends either directly or through +TIPs: Small spindles 2 (Ssp2) in Drosophila and

Slains in mammals (not shown; Al-Bassam and Chang, 2011; Currie et al., 2011; van der Vaart et al., 2012; Li et al., 2011; Lowery et al., 2010). Stathmins

(STMN) sequester tubulin (Manna et al., 2009), and doublecortin (DCX) binds MT plus ends independently (Bechstedt and Brouhard, 2012). (2) Actin dynamics.

The Shot–actin linkage is influenced by actin regulators (red stippled arrows) that coordinate the dynamics and structure of F-actin networks (cortex, lamellipodia,

filopodia; Pak et al., 2008; Xu et al., 2013). In the context of filopodia formation, Shot contributes to the regulation of F-actin through binding of Exba/eIF5C at its

EF-hand motifs (Lee et al., 2007; Sanchez-Soriano et al., 2009). (3) MT shaft stability. In addition, Shot binds along MT shafts and stabilises them independent of

its linkage to F-actin (Sanchez-Soriano et al., 2009; Alves-Silva et al., 2012). Other MT shaft binders are Tau which protects MTs against the MT-severing protein

Katanin in axons (not shown; Qiang et al., 2006), and mammalian MAP1B can cross-link MTs as well as mediate actin-MT linkage during axon growth (Bouquet

et al., 2007; Riederer, 2007). Tau, MAP1B and Shot (in an EB1-independent mode) influence MT polymerisation kinetics through yet unknown mechanisms

(stippled grey arrow; Alves-Silva et al., 2012; Feinstein and Wilson, 2005; Tymanskyj et al., 2012).

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might regulate the number of filopodia directly through its

function as an ABP, or indirectly through acting as a scaffold forRobo receptors, since loss of Robo has been previously shownto lead to an increase in the number of filopodia (Murray and

Whitington, 1999).

Use of the fly to study the functional integrationof ABPs during filopodia regulationDrosophila growth cones have been used to carry out loss-of-function analyses of other ABP-encoding genes, including Arpc1

(ARP2), Arp3 (ARP3), capping protein A (CAPZA), capping

protein B (CAPZB), chickadee (profilin), enabled (ENAH, also

known as VASP), singed (fascin) and tropomyosin 1 (TPM1)(human homologues are given in the brackets) (Goncalves-Pimentel et al., 2011; Kraft et al., 2006; Sanchez-Soriano et al.,

2010). All these data and tools can now be combined tounderstand the functional networks of ABPs, as is best illustratedfor studies of filopodia formation. Filopodia are prominent actin-

based structures, and data from numerous studies in a wide rangeof cellular models have led to the formulation of two compositemodels of filopodia formation (Mattila and Lappalainen, 2008).In the ‘convergent elongation model’, Arp2/3 generates actin

filaments in lamellipodia, and ENAH/VASP aggregates their plusends to give rise to elongating filopodial bundles. In the ‘de novo

nucleation model’, formin molecules aggregate to generate

clusters of new actin filaments, which then directly elongateinto parallel filopodial bundles.

Drosophila neurons were used to test whether these two

models co-exist in neurons. Single-mutant analyses of the factorsessential in these models (i.e. Enabled, the formin DAAM anddifferent Arp2/3 complex components) all caused a partial butsignificant reduction in filopodia numbers, in agreement with the

potential co-existence of both models (Goncalves-Pimentel et al.,2011; Matusek et al., 2008). When loss of the Arp2/3 componentSop2 was combined with functional loss of DAAM in the same

neurons (DAAM2/2 Sop22/2 double-mutant neurons), filopodiawere completely absent and F-actin levels were severely depleted(Goncalves-Pimentel et al., 2011). To our knowledge, a genetic

condition that depletes F-actin to this degree has not beenreported for other eukaryotic systems. It indicates that Arp2/3 andDAAM are the prevailing nucleators in embryonic Drosophila

neurons. The fact that the two nucleators enhance each others’

phenotypes indicates that they work in distinct complementaryfunctional pathways during filopodia formation. This isconsistent with the biochemical evidence that Arp2/3 and

formins nucleate actin filaments through different molecularmechanisms (Chesarone and Goode, 2009; Mattila andLappalainen, 2008). However, it does not yet resolve whether

these two nucleators contribute to two entirely separate processesof filopodia formation.

To make this distinction, genetic interaction studies usingpartial loss of gene functions are very helpful. For example, in

heterozygous gene constellation (i.e. one mutant and one normalgene copy), the levels of the proteins they encode tend to be onlymoderately reduced, and, typically, this reduction is not sufficient

to cause a phenotype. Accordingly, neurons that are heterozygousfor DAAM, Sop2 or ena show normal filopodia numbers(Goncalves-Pimentel et al., 2011). However, any combinations

of these heterozygous conditions (transheterozygous mutantneurons: DAAM2/+ Sop22/+ or DAAM2/+ ena2/+ or Sop22/+

ena2/+) become functionally rate-limiting and lead to a strong

reduction in filopodia numbers (Goncalves-Pimentel et al., 2011).

These findings are inconsistent with regard to whether twocompletely separate modes of filopodia formation exist, but dosuggest that the pathways downstream of the two nucleators

converge at some point. Therefore, these authors suggested amore general model of convergent elongation, in which not onlyArp2/3-derived but also DAAM-derived actin filaments can beused by Ena to be transformed into filopodial actin bundles

(Goncalves-Pimentel et al., 2011). These examples illustrate howsystematic genetic analyses used in Drosophila neurons cancontribute to the formulation of new mechanistic models of

cytoskeletal regulation.

Use of the fly to study mechanisms of actin–MTlinkage during axonal growth and maintenanceActin–MT crosstalk is an important aspect of cytoskeletalregulation in growing axons, but its mechanisms are littleunderstood (Box 2). Promising contributions have been made by

work on the above mentioned spectraplakins Shot and ACF7.Besides filopodia-associated phenotypes, we have shown thatloss of Shot in fly neurons and loss of the mouse homologueACF7 in mouse neurons result in another conserved phenotype –

disorganised axonal MT bundles that is coupled with reducedaxon growth (Sanchez-Soriano et al., 2009). Detailed work onShot has shown that these functions depend on actin–MT linkage.

Structure–function analyses have revealed that functions of Shotin MT organisation depend on three simultaneous molecularinteractions (Fig. 2): (1) binding of its N-terminal calponin

homology domains to F-actin, (2) association to MTs throughtwo C-terminal domains (the Gas2-related domain and thepositively charged C-terminal tail), and (iii) binding of C-

terminal tail to EB1 (end-binding protein 1) at MT plus ends(Alves-Silva et al., 2012; Bottenberg et al., 2009; Lee andKolodziej, 2002; Sanchez-Soriano et al., 2009). These findingssuggest a model in which Shot binds to the plus ends of

polymerising MTs and guides them along actin structures in thedirection of axon growth (Alves-Silva et al., 2012). This model isfurther supported by the observation that frequent off-track

extensions of MTs are seen during live imaging of shot mutantneurons; this can explain the observed MT disorganisation anddiminished axon growth (off-track MTs are less likely to

contribute to axon extension). The guidance model is alsosupported by shot-like MT-disorganisation phenotypes observedin EB1-deficient, as well as in shot2/+ eb12/+ transheterozygous

mutant neurons (Alves-Silva et al., 2012), and, furthermore, isconsistent with proposed roles of ACF7 in guiding MT extensionalong actin stress fibres towards focal adhesions in non-neuronalcells (Kodama et al., 2004).

Notably, the MT guidance model offers a promising molecularplatform for further research. On the one hand, it can be used tounderstand the roles of other cytoskeletal regulators during MT

guidance, for example of ABPs or the various proteins that bindand regulate MT plus ends (see details in Fig. 2). On the otherhand, the MT guidance model suggests a potential mechanismcontributing to the maintenance of axons in the mature nervous

system, as will be explained in the following.

The second mammalian homologue of Shot, dystonin, isstrongly expressed in peripheral neural ganglia (Leung et al.,

2001), which, in dystonin mutant mice, undergo a severeneurodegeneration at postnatal stages that is referred to asHSAN (hereditary sensory autonomic neuropathy) (Duchen et al.,

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1963; Duchen et al., 1964). The underlying mechanisms remain

inconclusive and were proposed to be associated with roles ofdystonin in intermediate filament organisation and axonaltransport, and in ER or Golgi dysfunction (Ferrier et al., 2013;

Young and Kothary, 2007). We propose instead that HSANpathology might be primarily caused by defects in MT guidanceand stabilisation on the basis of the following argument. AxonalMT bundles are believed to be stabilised through structural

MTBPs, such as tau or MAP1B (Fig. 2) (Chilton and Gordon-Weeks, 2007). At the same time, there appears to be a steadyand dynamic MT turnover, as suggested by continued MT

polymerisation events in mature axons (Kollins et al., 2009).Spectraplakins can regulate both functions through twofunctionally conserved domains at their C-terminus: the Gas2-

related domain can stabilise MTs against depolymerisation,whereas the C-terminal tail mediates the aforementioned bindingto EB1 required for MT guidance. These domains jointly mediate

a robust association along the shaft and at the tip of MTs that isrequired for both MT guidance and stabilisation (Fig. 2) (Alves-Silva et al., 2012; Honnappa et al., 2009; Sun et al., 2001).Absence of either domain in Shot causes severe MT

disorganisation and loss of MT stability in axons of developingfly neurons (Alves-Silva et al., 2012; Sanchez-Soriano et al.,2009) and, in addition, mature sensory neurons of dystonin

mutant mice, which are prone to degeneration display severe MTdisorganisation and loss of MT stability (Ferrier et al., 2013;Yang et al., 1999). Consistently, a recently reported frame-shift

mutation in the human dystonin gene functionally deletes theGas2-related domain and the C-terminal tail and causes a severeform of HSAN that mirrors the degenerative pathology of dystonin

mutant mice (Edvardson et al., 2012). These data suggest thatspectraplakins perform similar MT-regulating functions during thedevelopment of fly neurons and the maintenance of sensorymammalian neurons. Severe MT disorganisation and instability

caused by absence of spectraplakins can, therefore, be expected toaffect vital functions such as axonal transport and impactnegatively on neuronal survival (Sheng and Cai, 2012).

This example shows how functional studies of cytoskeletalregulators during axon growth in the fly can provide new ideasand testable hypotheses for research on other aspects of neuronalcytoskeleton, even in other animals.

Conclusions and future perspectivesIn this Commentary, we have discussed how systematic andcombinatorial genetic studies of different ABPs and MTBPs in

the neuronal system of the fly can improve our understandingof cytoskeletal regulation during a biological process such asaxon growth. Considering the high degree of evolutionary

conservation of actin, tubulin and their regulators, theenormous similarities of cytoskeletal dynamics in fly andvertebrate neurons, and the similarities of mutant phenotypes

reported so far, we feel that insights gained in the fly system willbe informative for cytoskeletal research in mammalian or othervertebrate neurons.

Understanding the cytoskeletal regulation that underpins

cellular behaviours will have important implications andapplications. For example, we still have little understanding ofhow signalling mechanisms instruct the cytoskeleton to influence

growth cone behaviours (Huber et al., 2003). Knowing how thevarious ABPs and MTBPs that are targeted by particularsignalling events contribute to the cellular cytoskeletal

machinery will greatly help to close this knowledge gap.

Understanding these cellular roles of cytoskeletal regulators isalso the first step on the long path to developing mathematicalmodels that can describe the cytoskeletal dynamics that are at the

basis of different cellular processes (Oelz et al., 2008), and it willfacilitate the introduction of systems approaches for cell biologyresearch (Liberali and Pelkmans, 2012). Furthermore, theprincipal strategies described here for axon growth can also be

applied to the investigation of the roles and regulation of thecytoskeleton in other relevant areas, such as synapse formation,axonal pruning and Wallerian degeneration, axon regeneration

and neurodegeneration. Notably, for all of these processes,suitable Drosophila models have already been established, thushopefully paving the way for rapid progress (Bossing et al., 2012;

Fang and Bonini, 2012; Gistelinck et al., 2012; Goellner andAberle, 2012; Jaiswal et al., 2012).

AcknowledgementsWe would like to thank Tom Millard, Laura Anne Lowery, PhillipGordon-Weeks, Joszef Mihaly and Erik Dent for their very helpfuland constructive comments on this manuscript.

FundingAuthors on this Commentary were funded by the Biotechnology andBiological Sciences Research Council (BBSRC) [grant numbers BB/D526561/1, BB/I002448/1]. Deposited in PMC for immediate release.

Supplementary material available online at

http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.126912/-/DC1

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OMIM entries of cytoskeletal genes

HUMAN NEURONAL DISORDERS

1. ACTB 7 (ACTIN, BETA; 102630) - Baraitser-Winter syndrome 2 243310 (including microcephaly, intellectual disability, seizures, and hearing loss); Dystonia, juvenile-onset 607371 (with neurodegenerative traits)

2. ACTG1 (ACTIN, GAMMA 1; 102560) - Baraitser-Winter syndrome 2 (microcephaly, intellectual disability, hearing loss, anterior-to-posterior gradient lissencephaly of the pachygyria or pachygyria-band type); 614583; Deafness, autosomal dominant 20/26 604717

3. APC (ADENOMATOUS POLYPOSIS OF THE COLON; 611731) - adenomatous polyposis coli 175100; Colorectal cancer, somatic 114500; Desmoid disease, hereditary 135290; Gastric cancer, somatic 613659; Hepatoblastoma, somatic 114550 ; Brain tumor-polyposis syndrome 2

4. CEP290 (CENTROSOMAL PROTEIN 290kDa; 610142) - localises to cilia and centrosome (ATP-binding, CC, tropomyosin-like domains); Leber congenital amaurosis 10 611755 (retinal degeneration and impaired olfaction)

5. CLIP2/CLIP115 (CAP-GLY domain containing linker protein 2; 603432) - Williams-Beuren syndrome, aortic stenosis with mental retardation (WBS; 194050) - supported by a mouse model.

6. DCTN1 (DYNACTIN 1; 601143) - Perry syndrome 168605; unusual neuropsychiatric disorder with mental depression and late onset of Parkinsonism; Neuropathy, distal hereditary motor, type VIIB 607641; susceptibility to Amyotrophic lateral sclerosis (neurodegenerative disorder) 105400

7. DCX (DOUBLECORTIN; 300121) - tubulin-binding, regulator of tubulin nucleation and polymerisation; Lissencephaly, X-linked 300067; X-linked subcortical laminal heteropia/double cortex (mild lissencephaly) 300067

8. DES (DESMIN; 125660) - type III intermediate filament; neurogenic Scapuloperoneal syndrome, Kaeser type (some individuals: mild demyelinating polyneuropathy and electromyography demonstrated acute and chronic denervation in both proximal and distal muscles) 181400

9. DIAPH3 (DIAPHANOUS, DROSOPHILA, HOMOLOG OF, 3; 614567) - Auditory neuropathy, autosomal dominant, 1 609129

10. DMD (DYSTROPHIN; 300377) - Becker muscular dystrophy 300376; Cardiomyopathy, dilated, 3B 302045; Duchenne muscular dystrophy accompanied by mental retardation 310200

11. DST/BPAG1/BP240 (dystonin; 113810) - Neuropathy, hereditary sensory and autonomic, type VI 614653

12. DYNC1H1 (CYTOPLASMIC DYNEIN HEAVY CHAIN; 600112) - Charcot-Marie-Tooth disease, axonal, type 20 614228; Mental retardation, autosomal dominant 13 614563; Spinal muscular atrophy, lower extremity, autosomal dominant 158600

13. FSCN2 (FASCIN, SEA URCHIN, HOMOLOG OF, 2; 607643) - Retinitis pigmentosa (retinal degeneration) 30 607921

14. ELP1 (IKBKAP; 603722) - tubulin acetylase; Dysautonomia, familial 223900

15. ESPN (ESPIN / ECTOPLASMIC SPECIALIZATION PROTEIN; 606351) - multifunctional actin-bundling protein; Deafness, autosomal recessive 36 609006; Deafness, neurosensory, without vestibular involvement, autosomal dominant

16. FLNA (FILAMIN A, ALPHA) - actin-binding; many skeletal disorders; periventricular Heterotopia (neuronal migration defect) 300049 and 300537; Intestinal pseudoobstruction, neuronal (including abnormal intermediate signal in the peritrigonal white matter) 300048

A. Prokop et al. - Understanding cytoskeletal machinery of neurons 2

17. GAN (GAN GENE; 605379) - putative cytoskeleton binding; Giant axonal neuropathy-1

256850

18. GFAP (GLIAL FIBRILLARY ACIDIC PROTEIN; 137780) - intermediate-filament (IF) protein that is highly specific for astroglia; Alexander disease (megalencephaly in infancy accompanied by progressive spasticity and dementia with most patients dying by 10 years) 203450

19. GSN (GELSOLIN; 137350) - Amyloidosis, Finnish type (including cranial and peripheral neuropathies) 105120

20. HDAC4 (HISTONE DEACETYLASE 4; 605314) - deacetylase, likely to act on tubulin; Brachydacytly-mental retardation syndrome 600430

21. INF2 (INVERTED FORMIN; 610982) - Charcot-Marie-Tooth disease, dominant intermediate E 614455; focal segmental glomerulosclerosis (renal disease) 613237

22. KATNAL2 (KATANIN P60 ATPASE-CONTAINING SUBUNIT A-LIKE 2; ) - autism spectrum diseases (reference)

23. KIAA0196 (STRUMPELLIN; 610657) - potential cytoskeletal regulator; Spastic paraplegia 8, autosomal dominant 603563

24. KIF1A (KINESIN FAMILY MEMBER 1A; 601255) - Mental retardation, autosomal dominant 9 614255; Neuropathy, hereditary sensory, type IIC 614213; Spastic paraplegia 30, autosomal recessive 610357

25. KIF1B (KINESIN FAMILY MEMBER 1B; 605995) - Charcot-Marie-Tooth disease, type 2A1 (hereditary motor and sensory neuropathies) 118210; Pheochromocytoma 171300; susceptibility to Neuroblastoma 1 256700; increased risk of nonsyndromic paraganglioma

26. KIF5A KINESIN or NKHC (KINESIN FAMILY MEMBER 5B, HEAVY CHAIN, NEURON-SPECIFIC; 602821) - Spastic paraplegia 10, autosomal dominant 604187

27. KIF7 (KINESIN FAMILY MEMBER 7; 611254) - Acrocallosal syndrome/Joubert syndrome 12 (mental retardation syndrome with brain abnormalities) 200990

28. KIF11 or KNSL1 or KHC (KINESIN FAMILY MEMBER 11, KINESIN-LIKE 1; KINESIN HEAVY CHAIN; 148760) - Microcephaly with or without chorioretinopathy, lymphedema, or mental retardation 152950

29. LIS1 (LISSENCEPHALY 1; 607432; official name: PLATELET-ACTIVATING FACTOR ACETYLHYDROLASE, ISOFORM 1B, ALPHA SUBUNIT; PAFAH1B1; 601545) - lissencephaly 1 607432; subcortical laminar heterotopia 607432

30. MAPT (MICROTUBULE-ASSOCIATED PROTEIN TAU; 157140) - Frontotemporal Dementia with or without parkinsonism (600274); Pick disease 172700; Supranuclear palsy, progressive 601104; Supranuclear palsy, progressive atypical 260540

31. MYH9 (MYOSIN IIA HEAVY CHAIN 9; 160775) - Deafness, autosomal dominant 17 603622; Epstein syndrome 153650; Fechtner syndrome 153640: Macrothrombocytopenia and progressive sensorineural deafness 600208; May-Hegglin anomaly 155100; Sebastian syndrome 605249

32. MYO1A (MYOSIN IA; 607841) - Deafness, autosomal dominant 48 607841

33. MYO5A (MYOSIN VA; MYO5; MYOXIN; MYOSIN HEAVY CHAIN 12; 160777) - Griscelli syndrome, type 1 214450 (hypomelanosis with a primary neurologic deficit including motor development delay and mental retardation)

34. MYO6 (MYOSIN VI, 600970) - nonsyndromic deafness (hearing loss without related signs and symptoms affecting other parts of the body) called DFNB37, likely disrupting stereocilia

35. MYO7A (MYOSIN VIIA; 276903) - Deafness, autosomal dominant 11 601317; Deafness, autosomal recessive 2 600060; Usher syndrome, type 1B (hearing loss or deafness and progressive vision loss) 276900

36. MYO15A (MYOSIN XVA; 602666) - Deafness, autosomal recessive 3 600316

A. Prokop et al. - Understanding cytoskeletal machinery of neurons 3

37. NDE1 (NUDE NUCLEAR DISTRIBUTION E HOMOLOG 1; 609449) - part of a multiprotein

complex that regulates dynein function; Lissencephaly 4 (with microcephaly) 614019

38. NEFH (NEUROFILAMENT, HEAVY POLYPEPTIDE; 162230) - susceptibility to Amyotrophic lateral sclerosis 105400

39. NEFL (NEUROFILAMENT, LIGHT POLYPEPTIDE; 162280) - Charcot-Marie-Tooth disease, type 1F 607734; Charcot-Marie-Tooth disease, type 2E 607684

40. PRPH (PERIPHERIN; 170710) - type III intermediate filament; susceptibility to Amyotrophic lateral sclerosis (neurodegenerative disorder characterized by the death of motor neurons) 105400

41. RDX (RADIXIN; 179410) - Deafness, autosomal recessive 24 611022 (autosomal recessive sensorineural deafness)

42. SPAST (SPASTIN; 604277) - Spastic paraplegia 4, autosomal dominant 182601

43. TUBA1A (ALPHA 1 TUBULIN) - Lissencephaly 3 611603

44. TUBA8 (ALPHA 8 TUBULIN) - Polymicrogyria with optic nerve hypoplasia 613180

45. TUBB3 (ß3 TUBULIN; 602661) - Fibrosis of extraocular muscles, congenital, 3A (600638); Cortical dysplasia, complex, with other brain malformations (CDCBM) 614039, a more severe disorder of aberrant neuronal migration and disturbed axonal guidance resulting in mental retardation.

46. TUBB2B (ß2 TUBULIN; 612850) - asymmetric Polymicrogyria 610031; malformation of cortical development characterized by an excessive number of small gyri with abnormal lamination

47. TRIOBP (TRIO and F-actin binding protein; 609761) - organization of actin filaments; Deafness, autosomal recessive 28 609823

48. SPAST (SPASTIN; 604277) - MT severer; Spastic paraplegia 4, autosomal dominant 182601

49. SPTAN1 (SPECTRIN, ALPHA, NONERYTHROCYTIC 1; 182810) - Epileptic encephalopathy, early infantile, 5 613477

50. SPTBN2 (SPECTRIN, BETA, NONERYTHROCYTIC, 2; 604985) - Spinocerebellar ataxia 5 600224

51. TUBGCP6 (TUBULIN-GAMMA COMPLEX-ASSOCIATED PROTEIN 6; 610053) - Microcephaly and chorioretinopathy with or without mental retardation 251270

ANIMAL MODELS OF NEURONAL DISORDERS (where no human brain disorder reported)

52. CFL1 (COFILIN 1; 601442) - deacetylase, likely to act on tubulin; mouse model with morphological abnormalities of cortex

53. DBN1 (DREBRIN E; 126660) - age-related significant decrease in postsynaptic drebrin in a transgenic mouse model

54. DPYSL2 or DRP2 or CRMP2 / COLLAPSIN RESPONSE MEDIATOR PROTEIN 2 (DIHYDROPYRIMIDINASE-LIKE 2; 602463) - tubulin dimer binding - rat model for inflammatory and neuropathic hypersensitivity

55. HDAC1 (HISTONE DEACETYLASE 1; 601241) - mouse model: Deletion of both Hdac1 and Hdac2 led to major abnormalities of cortical, hippocampal, and cerebellar development, whereas deletion of either individually had no apparent effect on neuronal development.

56. SPTBN4 (SPECTRIN, BETA, NONERYTHROCYTIC, 4; QUIVERING; 606214) - autosomal recessive mouse mutation 'quivering' (qv) produces progressive ataxia with hindlimb paralysis, deafness, and tremor

57. STMN1 (STATHMIN 1; 151442) - increased amounts of MTs in brain and behavioural deficits in mouse models (memory, parental care, fear)

A. Prokop et al. - Understanding cytoskeletal machinery of neurons 4

58. TBCE (TUBULIN-SPECIFIC CHAPERONE E; 604934) - Hypoparathyroidism-retardation-

dysmorphism syndrome (skeleton) 241410; Kenny-Caffey syndrome-1 (skeleton) 244460; mouse model: progressive caudiocranial motor axon degeneration from the age of 2 weeks

59. UTRN (UTROPHIN; 128240) - mice deficient for both dystrophin and utrophin pheoncopy DMD in humans, including severe progressive muscular dystrophy resulting in premature death, ultrastructural myotendinous and neuromuscular junction abnormalities

NON-NEURONAL HUMAN DISORDERS

60. ACTA1 (ACTIN, ALPHA 1, skeletal muscle; 102610) - different myopathies: Myopathy, actin, congenital, with excess of thin myofilaments 161800; Myopathy, congenital, with fiber-type disproportion 1 255310; Myopathy, nemaline, 3 161800

61. ACTA2 (ACTIN, ALPHA 2, smooth muscle, aorta; 102620) - Aortic aneurysm, familial thoracic 6 611788; Moyamoya disease 5 614042; Multisystemic smooth muscle dysfunction syndrome 613834

62. ACTC1 (ACTIN, ALPHA, CARDIAC MUSCLE; 102540) - Atrial septal defect 5 612794; Cardiomyopathy, dilated, 1R 613424; Cardiomyopathy, familial hypertrophic, 11 612098; Left ventricular noncompaction 4 613424

63. ACTN2 (ACTININ, ALPHA-2; 102573) - Cardiomyopathy, dilated, 1AA 612158

64. ACTN4 (ACTININ, ALPHA-4; 604638) - Glomerulosclerosis, focal segmental, 1 603278

65. {Hypertension, essential, salt-sensitive} 145500

66. ANK2 (ANKYRIN 2, neuronal) - heart disorders

67. BFSP2 (BEADED FILAMENT STRUCTURAL PROTEIN 2; 603212) - intermediate filament; Cataract, autosomal dominant, multiple types 1 611597; Cataract, congenital 604219; Cataract, juvenile-onset 604219

68. CFL2 (COFILIN 2; 601443) - Nemaline myopathy 7 (NEM7; 610687)

69. CTNNB1 (CATENIN, BETA-1; 116806) - Hepatocellular carcinoma 114550; Ovarian cancer 167000; Pilomatricoma 132600

70. DNAAF3 (DYNEIN, AXONEMAL, ASSEMBLY FACTOR 3; 614566) - Ciliary dyskinesia, primary, 2 606763

71. DNAH5 (DYNEIN, AXONEMAL, HEAVY CHAIN 5) - Ciliary dyskinesia, primary, 3, with or without situs inversus 608644 (chronic respiratory tract infections, abnormally positioned internal organs, infertility)

72. DNAH11 (DYNEIN, AXONEMAL, HEAVY CHAIN 11; 603339) - Ciliary dyskinesia, primary, 7, with or without situs inversus 611884

73. DNAI1 (DYNEIN, AXONEMAL, INTERMEDIATE CHAIN 1; 604366) - Ciliary dyskinesia, primary, 1, with or without situs inversus 244400

74. DNAI2 (DYNEIN, AXONEMAL, INTERMEDIATE CHAIN 2; 605483) - Ciliary dyskinesia, primary, 9, with or without situs inversus 612444

75. DSP (DESMOPLAKIN; 125647) - Arrhythmogenic right ventricular dysplasia 8 607450; Dilated cardiomyopathy with woolly hair and keratoderma 605676; Epidermolysis bullosa, lethal acantholytic 609638; Keratosis palmoplantaris striata II 612908; Skin fragility-woolly hair syndrome 607655

76. DYNC2H1 (DYNEIN, CYTOPLASMIC 2, HEAVY CHAIN 1; 603297) - skeletal and early developmental defects: Asphyxiating thoracic dystrophy 3 613091; Short rib-polydactyly syndrome, type II, digenic 263520; Short rib-polydactyly syndrome, type III 263510

77. FLNB (filamin B, beta; 603381) - actin-binding; skeletal disorders: Atelosteogenesis, type I 108720; Atelosteogenesis, type III 108721; Boomerang dysplasia 112310; Larsen syndrome 150250; Spondylocarpotarsal synostosis syndrome 272460

78. FLNC (FILAMIN C; FLN2; 102565) - Myopathy, distal, 4 614065; Myopathy, myofibrillar, 5 609524)

A. Prokop et al. - Understanding cytoskeletal machinery of neurons 5

79. GSN (GELSOLIN; 137350) - Amyloidosis, Finnish type 105120 (lattice corneal dystrophy type

II)

80. JUP (JUNCTION PLAKOGLOBIN, PKGB, GAMMA CATENIN, DESMOPLAKIN III; 173325) - Arrhythmogenic right ventricular dysplasia 12 611528; Naxos disease 601214

81. KIF21A (KINESIN FAMILY MEMBER 21A) - Fibrosis of extraocular muscles, congenital, 1 135700; Fibrosis of extraocular muscles, congenital, 3B 135700

82. KIF22 or KNSL4 or OBP (KINESIN FAMILY MEMBER 22, KINESIN-LIKE 4, ORIP-BINDING PROTEIN; 603213) - Spondyloepimetaphyseal dysplasia with joint laxity, type 2 603546; skeletal disorder without intellectual impairment

83. MYBPC1 (MYOSIN BINDING PROTEIN C, slow type; 160794) - Arthrogryposis, distal, type 1B 614335

84. MYH2/2A/S2A (MYOSIN, HEAVY CHAIN 2/2A/S2A, SKELETAL MUSCLE, ADULT; 160740) - Inclusion body myopathy-3 605637

85. MYH3 (MYOSIN, HEAVY CHAIN 3, skeletal muscle, embryonic; 160720) - Arthrogryposis, distal, type 2B (muscular) 601680

86. MYH7 (MYOSIN, HEAVY CHAIN 7, cardiac muscle, beta; 160760) - myopathies

87. MYH11 or SMMHC (MYOSIN, HEAVY CHAIN 11, SMOOTH MUSCLE; SMOOTH MUSCLE MYOSIN HEAVY CHAIN; 160745) - Aortic aneurysm, familial thoracic 4 132900

88. MYL2 or RLC (MYOSIN, LIGHT CHAIN 2, REGULATORY, CARDIAC, SLOW; REGULATORY LIGHT CHAIN OF MYOSIN; 160781) - Cardiomyopathy, familial hypertrophic, 10 608758

89. MYL3 or MLC1SB/1V (MYOSIN, LIGHT CHAIN 3, ALKALI, VENTRICULAR, SKELETAL, SLOW; MYOSIN, LIGHT CHAIN 1, SLOW, B/VENTRICULAR; 160790) - Cardiomyopathy, familial hypertrophic, 8 608751

90. NEM3 (NEMALINE MYOPATHY 3; 161800) - Myopathy, nemaline, 3 161800; Myopathy, actin, congenital, with excess of thin myofilaments 161800

91. NF2 (NEUROFIBROMIN 2, merlin/Moesin-Ezrin-Radixin-Like Protein; 607379) - Meningioma, NF2-related, somatic 607174; Neurofibromatosis, type 2 101000; Schwannomatosis 162091

92. PCNT (PERICENTRIN; 605925) - Microcephalic osteodysplastic primordial dwarfism, type II 210720

93. PLEC1 (PLECTIN 1; 601282) - Epidermolysis bullosa simplex with pyloric atresia 612138; Epidermolysis bullosa simplex, Ogna type 131950; Muscular dystrophy with epidermolysis bullosa simplex 226670; Muscular dystrophy, limb-girdle, type 2Q 613723

94. PKP1 (PLAKOPHILIN 1; 601975) - Ectodermal dysplasia/skin fragility syndrome 604536

95. PKP2 (PLAKOPHILIN 2; 602861) - Arrhythmogenic right ventricular dysplasia 9 609040

96. SPTA1 (SPECTRIN, ALPHA, ERYTHROCYTIC 1; 182860) - Elliptocytosis-2 130600; Pyropoikilocytosis 266140; Spherocytosis, type 3 270970

97. TNNI2 (TROPONIN I, FAST-TWITCH SKELETAL MUSCLE ISOFORM; 191043) - binds to actin in thin myofilaments; Arthrogryposis multiplex congenital, distal, type 2B 601680

98. TNNI3 (TROPONIN I, CARDIAC; 191044) - binds to actin in thin myofilaments; Cardiomyopathy, dilated, 1FF 613286; Cardiomyopathy, dilated, 2A 611880; Cardiomyopathy, familial hypertrophic, 7 613690; Cardiomyopathy, familial restrictive 115210

99. TPM1 (TROPOMYOSIN 1; 191010) - Cardiomyopathy, dilated, 1Y 611878; Cardiomyopathy, familial hypertrophic, 3 115196

100. TPM2 (TROPOMYOSIN 2; 190990) - Arthrogryposis multiplex congenita, distal, type 1/2B 108120 601680; Nemaline myopathy 609285

101. TPM3 (TROPOMYOSIN 3; 191030) - CAP myopathy 609284; Myopathy, congenital, with fiber-type disproportion 255310; Nemaline myopathy 1, autosomal dominant 609284

102. VCL (VINCULIN; 193065) - Cardiomyopathy, dilated, 1W 611407; Cardiomyopathy, familial hypertrophic, 15 613255

A. Prokop et al. - Understanding cytoskeletal machinery of neurons 6

103. WAS(P) (WISKOTT-ALDRICH SYNDROME PROTEIN; 300392) - various immuodeficiencies:

Neutropenia, severe congenital, X-linked 300299; Thrombocytopenia, X-linked 313900; Thrombocytopenia, X-linked, intermittent 313900; Wiskott-Aldrich syndrome 301000

NO REPORTED HUMAN DISORDER LINKS

104. ACTL6B (ACTIN-LIKE 6B; ACTIN-RELATED PROTEIN, ALPHA; 612458)

105. ACTIN, KAPPA (611266)

106. ACTN1 (ACTININ, ALPHA-1; 102575)

107. ACTN3 (ACTININ, ALPHA-3; 102574) - potential link to Sprinting performance

108. ACTR2 (ARP2; 604221) - Arp2/3 complex component

109. ACTR3 (ARP3; 604222) - Arp2/3 complex component

110. ADD2 (ADDUCIN 2; 102681)

111. ADD3 (ADDUCIN 3; 601568)

112. ARPC1A (ARP2/3 COMPLEX, SUBUNIT 1A; SOP2L; 604220)

113. ARPC1B (ARP2/3 COMPLEX, SUBUNIT 1B; ARC41; 604223)

114. ARPC2 (ARP2/3 COMPLEX, SUBUNIT 2; ARC34; 604224)

115. ARPC3 (ARP2/3 COMPLEX, SUBUNIT 3; ARC21; 604225)

116. ARPC4 (ARP2/3 COMPLEX, SUBUNIT 4; ARC20; 604226)

117. ARPC5 (ARP2/3 COMPLEX, SUBUNIT 5; ARC16; 604227)

118. CAMSAP1 (CALMODULIN-REGULATED SPECTRIN-ASSOCIATED PROTEIN 1; Ssp4; 613774) - MT minus end stabiliser

119. CAPG (CAPPING PROTEIN, GELSOLIN-LIKE; 153615)

120. CAPZA1 (CAPPING PROTEIN, ALPHA-1; 601580)

121. CAPZA2 (CAPPING PROTEIN, ALPHA-2; 601571)

122. CAPZA3 (CAPPING PROTEIN, ALPHA-3; 608722)

123. CAPZB (CAPPING PROTEIN, BETA; 601572)

124. CKAP5 (CYTOSKELETON-ASSOCIATED PROTEIN 5, CHTOG, XMAP215; 611142)

125. CLASP1 (CYTOPLASMIC LINKER-ASSOCIATED PROTEIN 1; 605852)

126. CLASP2 (605853)

127. CLIP1/CLIP170/RESTIN/RSN (179838) - weak links to Hodgkin disease/lymphoma

128. CORO1A (CORONIN 1A; 605000) - actin binding; mouse models with lymphocyte migration defects

129. CORO1B (CORONIN 1B; 609849) - actin binding

130. CORO1C (CORONIN 1C; 605269) - actin binding

131. CORO2A (CORONIN 2A; 602159) - actin binding

132. CORO2B (CORONIN 2B; 609849) - actin binding

133. CORO7 (CORONIN 7; 611668) - actin binding

134. CTTN (CORTACTIN; 164765) - actin binder and regulator

135. DAAM1 (DISHEVELLED-ASSOCIATED ACTIVATOR OF MORPHOGENESIS 1; 606626)

136. DAAM2 (DISHEVELLED-ASSOCIATED ACTIVATOR OF MORPHOGENESIS 2; 606627)

137. DBNL (DREBRIN-LIKE; 610106)

138. DNAH6 (DYNEIN, AXONEMAL, HEAVY CHAIN 6; 603336)

139. DNAH9 (DYNEIN, AXONEMAL, HEAVY CHAIN 9; 603330)

140. DNAL1 (DYNEIN, AXONEMAL, LIGHT CHAIN 1; 610062) - Ciliary dyskinesia, primary, 16 614017

141. DNALI1 (DYNEIN, AXONEMAL, LIGHT INTERMEDIATE POLYPEPTIDE 1; 602135)

A. Prokop et al. - Understanding cytoskeletal machinery of neurons 7

142. DSTN (DESTRIN/ACTIN DEPOLYMERIZING FACTOR; ADF; 609114) - mouse model with

irregular thickening of the corneal epithelium

143. DYNC1I2 (DYNEIN, CYTOPLASMIC 1, INTERMEDIATE CHAIN 2; 603331)

144. DYNC1I1 (DYNEIN, CYTOPLASMIC 1, INTERMEDIATE CHAIN 1; 603772)

145. DYNC1LI2 (DYNEIN, CYTOPLASMIC 1, LIGHT INTERMEDIATE CHAIN 2; 611406)

146. DYNLL1 (DYNEIN, LIGHT CHAIN, LC8 TYPE, 1; 601562)

147. DYNLL2 (DYNEIN, LIGHT CHAIN, LC8 TYPE, 2; 608942)

148. DYNLRB1 (DYNEIN, LIGHT CHAIN, ROADBLOCK TYPE, 1; 607167)

149. DYNLRB2 (DYNEIN, LIGHT CHAIN, ROADBLOCK TYPE, 2; 607168)

150. DYNLT1 (DYNEIN, LIGHT CHAIN, TCTEX TYPE, 1; 601554)

151. DYNLT3 (DYNEIN, LIGHT CHAIN, TCTEX TYPE, 3; 300302)

152. EB1 (MAPRE1/MICROTUBULE-ASSOCIATED PROTEIN, RP/EB FAMILY, MEMBER 1; 603108)

153. EB2 (MAPRE2; 605789)

154. EB3 (MAPRE3; 605788)

155. ELP3 (ELONGATION PROTEIN 3, S. CEREVISIAE, HOMOLOG OF; 612722) - tubulin acetylase

156. ENAH (ENAH enabled homolog; 609061)

157. EVPL (ENVOPLAKIN; 601590)

158. EZRIN/VIL2 (VILLIN 2; 123900) - mouse model with intestinal epithelial defects

159. FSCN1 (FASCIN, SEA URCHIN, HOMOLOG OF, 1; 602689)

160. HDAC5 (HISTONE DEACETYLASE 5; 300269) - likely tubulin deacetylase

161. HDAC8 (HISTONE DEACETYLASE 8; 300269) - likely tubulin deacetylase

162. PHF21A (PHD FINGER PROTEIN 21A/BRAF35/KIAA1696; 605315) - HDAC COMPLEX, 80-KD SUBUNIT, likely tubulin deacetylase

163. IQGAP1 (IQ MOTIF-CONTAINING GTPase-ACTIVATING PROTEIN 1; 603379)

164. IQGAP2 (IQ MOTIF-CONTAINING GTPase-ACTIVATING PROTEIN 2; 605401)

165. KATNA1 (KATANIN, p60 SUBUNIT, A1; 606696)

166. KATNAL1 (KATANIN p60 SUBUNIT A-LIKE 1; 614764)

167. KATNAL2 (KATANIN, p60 SUBUNIT, A-LIKE PROTEIN 2; 614697)

168. KATNB1 (KATANIN, p80 SUBUNIT, B1; 602703)

169. KIF2C or MCAK or KNSL6 (KINESIN FAMILY MEMBER 2C, MITOTIC CENTROMERE-ASSOCIATED KINESIN, KINESIN-LIKE 6; 604538) - +TIP and inhibitor of MT polymerisation

170. KIF5B (KINESIN FAMILY MEMBER 5B; 602809) - kif5B-/- mice were embryonic lethal with a severe growth retardation at 9.5 to 11.5 days postcoitum

171. KIF5C or NKHC2 (KINESIN FAMILY MEMBER 5C, NEURON-SPECIFIC KINESIN HEAVY CHAIN 2; 604593)

172. KIF20A or RABKINESIN 6 (KINESIN FAMILY MEMBER 20A; 605664) - frequently deleted in myeloid leukemias

173. KIF23 or KNSL5 or MKLP1 (KINESIN FAMILY MEMBER 23, KINESIN-LIKE 5; MITOTIC KINESIN-LIKE 1; 605064)

174. KLC1 or KLC or KNS2 (KINESIN LIGHT CHAIN 1, KINESIN 2; 600025) - mouse Klc1 predominantly in central and peripheral neuronal tissues

175. MACF1 (MICROTUBULE-ACTIN CROSS-LINKING FACTOR 1; 608271)

176. MAP1A (MICROTUBULE-ASSOCIATED PROTEIN 1A; 600178)

177. MAP1B (MICROTUBULE-ASSOCIATED PROTEIN 1B; 157129)

A. Prokop et al. - Understanding cytoskeletal machinery of neurons 8

178. MAP2 (MICROTUBULE-ASSOCIATED PROTEIN 2; 157130) - some indirect effect in Jnk1

(MAPK8; 601158)-null mouse

179. MSN (MOESIN; 309845) - actin-binding

180. MYH10 (MYOSIN, HEAVY CHAIN 10, NONMUSCLE; 160776) - homozygous null mice have 70% fewer, but larger, myocytes

181. MYL4 (MYOSIN, LIGHT CHAIN 4, ALKALI, ATRIAL, EMBRYONIC; 160770)

182. NINL (NINEIN-LIKE PROTEIN; 609580)

183. PARD3 (PARTITIONING-DEFECTIVE PROTEIN 3; 606745) - MT binding in neuronal polarisation

184. PFN1 (PROFILIN 1; 176610) - mouse model with immune response deficits

185. PFN2 (PROFILIN 2; 176590)

186. PFN2 (PROFILIN 2; 612812)

187. PLS1 (PLASTIN; FIMBRIN, 602734)

188. PKP3 (PLAKOPHILIN 3; 605561)

189. PKP4 (PLAKOPHILIN 4; 604276)

190. PPL (PERIPLAKIN; 602871)

191. PPP1R9A (PROTEIN PHOSPHATASE 1, REGULATORY SUBUNIT 9A; SPINOPHILIN; NEURABIN I; 602468) - actin binding

192. SPTB (SPECTRIN, BETA, ERYTHROCYTIC; 182870) - Anemia, neonatal hemolytic, fatal and near-fatal; Elliptocytosis-3; Spherocytosis, type 2

193. SPTBN1 (SPECTRIN, BETA, NONERYTHROCYTIC, 1; 182790)

194. SPTBN5 (SPECTRIN, BETA, NONERYTHROCYTIC, 5; 605916)

195. STMN2 or SCGN10 / SUPERIOR CERVICAL GANGLIA, NEURAL SPECIFIC 10 (STATHMIN-LIKE 2; 600621) - upregulated in familial dysautonomia (Cheishvili et al., 2011, Hum Mol Genet 20, 1585ff.)

196. STMN3 (STATHMIN-LIKE 3; 608362) - tubulin sequestering

197. TLN1 (TALIN 1; 186745)

198. TLN2 (TALIN 2; 607349)

199. TMOD1 (TROPOMODULIN; 190930)

200. TMOD2 (TROPOMODULIN 2; 602928)

201. TMOD3 (TROPOMODULIN 3; 605112)

202. TMOD4 (TROPOMODULIN 4; 605834)

203. TMSB4X (THYMOSIN, BETA-4, X CHROMOSOME; 300159) - actin monomer sequestering protein

204. TNNI1 (TROPONIN I, SLOW-TWITCH SKELETAL MUSCLE ISOFORM; 191042) - binds to actin in thin myofilaments

205. TPM4 (TROPOMYOSIN 4; 600317)

206. TUBE1 (TUBULIN, EPSILON-1; 607345)

207. TUBG1 (TUBULIN, GAMMA-1; 191135)

208. TUBG2 (TUBULIN, GAMMA-2; 605785)

209. TUBGCP4 (TUBULIN-GAMMA COMPLEX-ASSOCIATED PROTEIN 4; 609610)

210. TUBGCP5 (TUBULIN-GAMMA COMPLEX-ASSOCIATED PROTEIN 5; 608147)

211. TWF1 (TWINFILIN, DROSOPHILA, HOMOLOG OF, 1; 610932) - G-actin binding

212. associated with sensory neuropathy (SPTLC1, SPTLC2, WNK1, IKBKAP, NTRK1, NGFB, DNMT1

213. VASP (VASODILATOR-STIMULATED PHOSPHOPROTEIN; 601703)

214. WASL (WISKOTT-ALDRICH SYNDROME GENE-LIKE; 605056)

A. Prokop et al. - Understanding cytoskeletal machinery of neurons 9

215. WASF1 (WISKOTT-ALDRICH SYNDROME PROTEIN FAMILY, MEMBER 1, WAVE1,

SCAR1; 605035)

216. WASF2 (WISKOTT-ALDRICH SYNDROME PROTEIN FAMILY, MEMBER 2, WAVE2, SCAR2; 605875)

217. WASF3 (WISKOTT-ALDRICH SYNDROME PROTEIN FAMILY, MEMBER 3, WAVE3, SCAR3; 605068)