The organization and function of eukaryotic cells rely on the action of many different molecular motor proteins. Cytoplasmic dynein drives the movement of a wide range of cargoes towards the minus ends of microtubules, and these events are needed, not just at the single-cell level, but are vital for correct development. In the present paper, I review recent progress on understanding dynein's mechanochemistry, how it is regulated and how it binds to such a plethora of cargoes. The importance of a number of accessory factors in these processes is discussed.

Introduction

Cytoplasmic dynein-1 is a fascinating, complex molecular motor that drives the movement of an extraordinary range of cargoes towards the minus ends of microtubules. In a fibroblast, that means transport towards the cell centre, where the MTOC (microtubule-organizing centre) resides next to the nucleus. However, in other situations, such as polarized epithelial cells and neuronal dendrites, microtubules may be oriented with their plus ends towards the nucleus or have mixed polarity. Cytoplasmic dynein-1 (referred to herein as dynein) is a member of a family that includes many different axonemal (ciliary or flagellar) motors, and cytoplasmic dynein-2, whose primary role is in axoneme assembly [13].

The most obvious role for dynein is to transport discrete structures along microtubules. In animals, these range from individual proteins/protein complexes, through mRNPs (messenger ribonucleoproteins), to membranous organelles and chromosomes. However, dynein also drives the sliding of microtubules compared with each other, such as within the mitotic spindle, and of intermediate filaments along microtubules. Less obviously, dynein can be attached to a ‘fixed’ structure such as the cell cortex, where it can then pull on microtubules, and so playing a vital role in reorganizing the cell during migration or in positioning the mitotic spindle.

Dynein function is essential in animals, primarily because cells without functional dynein cannot divide properly. Interestingly, plants do not have dynein [3], and instead use multiple minus-end-directed kinesins for spindle assembly and cell division. Manipulating dynein function in organisms such as Drosophila and Caenorhabditis elegans has shown that it plays many different roles in development, particularly in the brain. For example, the branching of dendrites is dependent on dynein-driven transport of a range of cargoes [47], while neuronal survival depends on active retrograde transport of survival factors from the axon tip to the cell body [8]. Brain development is also crucially dependent on neuronal migration, which requires dynein to drive cell body and nuclear movement [911]. Even very subtle mutations in dynein subunits can lead to neurological defects [12,13]. Dynein's functions are simpler in fungi, where it pulls the nucleus into the daughter cell during budding yeast cell division, determines nuclear position within the hyphae of filamentous fungi and facilitates meiotic chromosome segregation [9]. While dynein transports membrane cargoes in filamentous fungi as it does in animal cells, yeast and plants use myosins for the same job.

Dynein composition and mechanism

Dynein is a 1.6 MDa complex containing two copies of the ATPase motor domain [heavy chain, DHC (dynein heavy chain)] and several additional subunits (Figure 1A) [1,3]. Two ICs (intermediate chains) bind directly to DHC, and then three different LCs (light chains), Tctex1, LC8 and LC7/roadblock, bind to the IC at separate sites. Two LICs (light intermediate chains) bind independently to the DHC. The ICs, LICs and LCs are each encoded by two genes in vertebrates, and the ICs and LICs may be alternatively spliced and present in different phosphoisoforms [1,3]. This variation suggests that different versions of dynein may perform distinct tasks by binding preferentially to specific cargo. While the ICs and LICs are only found within the dynein complex, the LCs have a variety of other interaction partners [1], complicating investigation of their function.

Dynein subunit composition, accessory proteins and proposed structure

Figure 1
Dynein subunit composition, accessory proteins and proposed structure

(A) Cartoon view of dynein and its subunits. The individual AAA domains are numbered. Interactions between accessory proteins and dynein subunits are indicated. (B and C) Proposed organization of a single DHC in the absence (B) and presence (C) of ATP. This model is based on X-ray crystallographic and EM data [1618]. The microtubule-binding domain and stalk extend from between AAA4 and AAA5, whereas the ‘strut’ (orange coil) extends from AAA5. The position of the linker (grey bar) is known [1618], whereas the position of the tail (grey wedge) is hypothetical. Other dynein subunits are not shown.

Figure 1
Dynein subunit composition, accessory proteins and proposed structure

(A) Cartoon view of dynein and its subunits. The individual AAA domains are numbered. Interactions between accessory proteins and dynein subunits are indicated. (B and C) Proposed organization of a single DHC in the absence (B) and presence (C) of ATP. This model is based on X-ray crystallographic and EM data [1618]. The microtubule-binding domain and stalk extend from between AAA4 and AAA5, whereas the ‘strut’ (orange coil) extends from AAA5. The position of the linker (grey bar) is known [1618], whereas the position of the tail (grey wedge) is hypothetical. Other dynein subunits are not shown.

DHC is an unusual member of the hexameric AAA+ (ATPase associated with various cellular activities) family of ATPases, since its six AAA domains are encoded by a single polypeptide. Hydrolysis of ATP by AAA1 and AAA3 is most important for motility, while hydrolysis by AAA2 and AAA4 may be regulatory, and AAA5 and AAA6 do not bind ATP [14]. DHC binds to microtubules via a small globular domain at the end of a 10 nm anti-parallel coiled coil, the stalk, that extends from between AAA4 and AAA5 [1517], with the base of the stalk being supported by a ‘buttress’ or ‘strut’ extending out of AAA5 [17,18]. There must be long-range communication across the head, since microtubule binding stimulates AAA1's ATPase activity, and microtubule binding is in turn affected by the nucleotide state of AAA1 [18] on the opposite side of the motor domain (Figures 1B and 1C). Force is thought to be generated by a large shift in position of a linker domain that joins the tail and AAA1 (Figures 1B and 1C) [1619]. In contrast, the angle of the stalk compared with the head remains fairly constant during the ATPase cycle [1619]. Surprisingly, a mutation in the tail domain reduces dynein's processivity and alters several other mechanochemical properties, suggesting that the tail plays an important role in movement, perhaps by co-ordinating the action of the two heads [20].

Studies on dynein's movement using purified protein have shown that it mainly takes steps of 8 nm (the distance between tubulin dimers), but occasionally steps of 16, 24 or 36 nm along the microtubule lattice [2123]. This intriguing behaviour is different from that of kinesins and myosins, which have one step size, and has also been seen in living cells [21]. Interestingly, increasing the load on the motor led to smaller steps, suggesting that dynein can ‘change gears’ depending on the force it has to generate [23]. As yet, the significance of this behaviour for the cell is unknown. Is the ability to alter step size fundamental to dynein's function, and is it a focus for regulation in vivo?

A conundrum is why dynein moves up to 10 times faster in living cells than in vitro (e.g. [24,25]). One possibility is simply that native cargoes might engage multiple active dyneins, with their speed of movement increasing by ~1 μm/s per motor recruited [25]. However, other studies have shown that a few (perhaps only one) dyneins are sufficient to drive rapid movement [2628], and that multiple motors may actually slow down translocation in vivo [29,30]. Nevertheless, the presence of more than one dynein on a cargo will certainly provide additional force to drive motility through the crowded cytoplasm. Additionally, it should also help cargoes side-step obstructions by switching to another microtubule [31]. The ability of dynein to take backward steps [22,23,32,33] is also likely to help in this regard.

Dynein accessory factors: effect on dynein activity

It is clear from many studies that dynein needs other molecules to work optimally. So far, however, none of the known dynein accessory proteins have been found to increase dynein's rate of translocation, but they have a range of effects on dynein's mechanochemistry, as well as being implicated in cargo binding (see below).

The dynactin complex (Figure 1A) [34] is needed for virtually all dynein functions, and enhances dynein's processivity (the number of steps a motor takes along the microtubule before falling off) [24,35,36], without affecting its rate of movement or its ability to step backwards [24,35,36]. This was thought to be due to the ability of the p150 subunit of dynactin to bind to microtubules either via its CAP-Gly (cytoskeleton-associated protein glycine-rich) domain or a neighbouring basic domain [35,37,38], thus serving to anchor dynein to the microtubule if both motor domains became unattached. Surprisingly, however, replacing p150 with a truncated version that could not bind microtubules had no effect on the processive movement of peroxisomes or mRNPs in vivo, nor on Golgi apparatus morphology [39,40]. Similar short p150 isoforms exist naturally [39], and yeast dynactin complexes containing truncated p150 still enhance dynein's processivity [41], suggesting that it is the p150–IC interaction that is important for stimulating dynein activity. Indeed, while an antibody to p150 that inhibits microtubule binding but not the IC–p150 interaction reduced the number of vesicles that associated with microtubules in vitro, it actually increased processive movement [27]. p150's microtubule binding activity may be necessary, however, when large forces are required, such as during nuclear migration [41], and for organizing microtubules within the cell [40].

An important development has been the identification of several proteins, such as NudF (nuclear distribution protein F)/LIS1 (lissencephaly 1), NudE (nuclear distribution protein E)/Nde1, NudE-like/Ndel1, BicD (Bicaudal D) and ZW10 (Zeste White 10), that are required for many dynein functions (reviewed in [14]). BicD and ZW10 are involved primarily in recruiting dynein to cargoes, and are discussed in the next section. LIS1 is mutated in lissencephaly, or smooth brain, where defective neuronal migration leads to disorganized brain architecture [11]. LIS1, Nde1 and Ndel1 engage in a complex network of interactions with each other, and with dynein/dynactin (Figure 1A), but are lost from purified dynein (e.g. [42,43]). LIS1 and the C-terminal domain of Ndel1 bind to the catalytic AAA1 domain of DHC [44,45], whereas Nde1 does not [43]; instead, Nde1 binds to IC and LC8 [14,46]. Interestingly, both Nde1 and Ndel1 can also bind to dynein via their N-terminal coiled-coil domains [47], which contain independent binding sites for IC [48] and LIS1 [47]. This has led to a model where the coiled-coil region of the Ndel1 dimer spans the dynein molecule from the IC in the cargo-binding domain to the head, where it interacts with LIS1 that is bound to the AAA1 domain of DHC [47]. This scaffolding role is proposed to be regulated by phosphorylation of the C-terminal of Ndel1 [47].

As one might expect from the location of these interactions, LIS1 and Nde1/Ndel1 have a profound influence on dynein's motor activity [42,43,49]. Rather surprisingly, however, LIS1 and Ndel1 individually decreased dynein's motor activity, which was restored when both LIS1 and Ndel1 were added together [42]. Subsequently, LIS1 has been shown to bind to dynein in its pre-power stroke conformation, enhancing its binding to microtubules and the force generated, but reducing the rate of movement due to increased pausing [43]. The presence of Nde1 promoted tight binding of LIS1 to dynein in any nucleotide state, and led to increased force generation [43]. However, Nde1 on its own reduced dynein's ability to bind to microtubules and to generate force [43]. Another study dissected the roles of different regions of Ndel1, and found that the C-terminus of Ndel1 was sufficient to cause dynein's release from microtubules, suggesting that it is a negative regulator [49]. Importantly, the binding of LIS1 by Ndel1 amino acids 44–183 was enough to reactivate dynein motility [49], and this truncation lacks the ability to bind to IC [48,49].

Altogether, these results highlight the crucial importance of dynein's accessory factors in regulating dynein mechanochemistry. An interesting question for the future is whether these interactors are a focus for regulating dynein function in the cell. For example, there is evidence that both the IC–p150 and Ndel1–DHC/LIS1 interactions are regulated by phosphorylation [47,50]. The function of dynein accessory proteins is not limited to influencing dynein activity; however, they also play roles in dynein–cargo interactions.

Targeting dynein to different cargoes

How dynein binds to its wide range of cargoes is a poorly understood topic. Given that there is considerable subunit heterogeneity, one appealing idea is that cargo-specific dyneins are defined by their composition of particular subunit splice variants or phosphoisoforms. Indeed, a wide range of IC splice forms are found in neuronal tissues [51], and the IC1B isoform specifically associates with TrkB (tropomyosin receptor kinase B)-positive endosomes in neurons [52]. However, only one splice form, IC2C, is found in non-neuronal mouse tissues [51].

Further evidence, both for and against the hypothesis that specific subunits define functional dynein subsets, has come from studies on the LIC. There are two LIC genes in vertebrates, but only one type of LIC is found in each dynein molecule, generating distinct dynein subtypes [53]. There are two clear instances of isoform-specific interactions: LIC1 with pericentrin [53] and LIC2 with the polarity molecule Par3 [54]. Melanosomes also possess only one of three possible LIC forms [55]. RNAi (RNA interference) knockdown of LICs individually has generated conflicting data over their roles in the secretory and endocytic pathways [2,56]. My group's work has shown that knockdown of both LICs always gives a more profound phenotype than single deletions, and that either LIC can rescue defects in organelle positioning in both the secretory and endocytic pathways (C. Villemant, A. Mironov, N. Flores-Rodriguez, P.G. Woodman and V.J. Allan, unpublished work). Similarly, both LICs influence the position of recycling endosomes and interact with the recycling endosome protein FIP3 (Rab11 family-interacting protein 3) (Table 1; [57,58]). In mitosis, single depletion of LIC1, but not LIC2, delayed mitotic progression [59], but we have again found that depleting both LICs causes more severe inhibition than LIC1 depletion alone (C. Villemant, A. Mironov, N. Flores-Rodriguez, P.G. Woodman and V.J. Allan, unpublished work).

Table 1
Selected dynein–cargo interactions

A range of recent and key earlier examples of dynein–cargo interactions are listed, along with the small GTPase involved, where appropriate. ASFV, African swine fever virus; ERGIC, endoplasmic recticulum–Golgi intermediate compartment; GKAP, guanylate kinase-associated protein; KIBRA, kidney- and brain-expressed protein.

Dynein subunit Cargo GTPase Other adaptor molecules Reference(s) 
Not specified Secretory vesicles, ERGIC, Golgi Rab6 BicD1/2 [77,79
Not specified Rab6-positive vesicles Rab6 BicD-related protein 1 [103
IC Lysosomes/autophagosomes in neurons  Snapin [112
IC Adenovirus particles  Hexon subunit [113
IC Kinetochores  ZW10 [61
LIC1 Endosomes Rab4  [114
LIC1 and LIC2 Endosomes Rab11a FIP3 [57,58
LIC1 Centrosomes  Pericentrin [53
LIC2 Cell cortex  Par3 [54
LIC Nuclei  Zyg12 [83
LIC APP (amyloid precursor protein)-containing vesicles in neurons  Unc-16/JIP3 [105
LIC Adenovirus particles  Hexon subunit [113
LC8 Cell cortex Cdc42 (cell division cycle 42; indirect) hDlg1/SAP97 via GKAP [115
LC8 RNPs (ribonucleoproteins)  Egalitarian (that binds BicD) [116
LC8 Piccolo–Bassoon transport vesicles (neurons)  Bassoon [117
LC8 Early endosomes  KIBRA, which binds sorting nexin 4 [118
LC8 ASFV  ASFV p54 [119
LC7/rbl Rab6 vesicles Rab6  [120
Dynein subunit Cargo GTPase Other adaptor molecules Reference(s) 
Not specified Secretory vesicles, ERGIC, Golgi Rab6 BicD1/2 [77,79
Not specified Rab6-positive vesicles Rab6 BicD-related protein 1 [103
IC Lysosomes/autophagosomes in neurons  Snapin [112
IC Adenovirus particles  Hexon subunit [113
IC Kinetochores  ZW10 [61
LIC1 Endosomes Rab4  [114
LIC1 and LIC2 Endosomes Rab11a FIP3 [57,58
LIC1 Centrosomes  Pericentrin [53
LIC2 Cell cortex  Par3 [54
LIC Nuclei  Zyg12 [83
LIC APP (amyloid precursor protein)-containing vesicles in neurons  Unc-16/JIP3 [105
LIC Adenovirus particles  Hexon subunit [113
LC8 Cell cortex Cdc42 (cell division cycle 42; indirect) hDlg1/SAP97 via GKAP [115
LC8 RNPs (ribonucleoproteins)  Egalitarian (that binds BicD) [116
LC8 Piccolo–Bassoon transport vesicles (neurons)  Bassoon [117
LC8 Early endosomes  KIBRA, which binds sorting nexin 4 [118
LC8 ASFV  ASFV p54 [119
LC7/rbl Rab6 vesicles Rab6  [120

The identification of LIC-interacting partners demonstrates that dynein can bind directly to cargo-specific components. Other direct interactions may involve the ICs or LCs (Table 1). Confusingly, some cargoes have several potential dynein ‘receptors’, while others have none as yet. Since the LCs also function outside the dynein complex, it is particularly important that LC interactors are shown to recruit the dynein whole complex, not just the LC, but this has not always been done.

Long before any of these cargo-specific dynein-binding partners were identified, dynactin was proposed to recruit dynein to structures ranging from kinetochores [60] to membranes [34]. In mitosis, dynein is needed at the kinetochore both to drive the movement of chromosomes along microtubules during chromosome alignment, and to remove components of the SAC (spindle assembly checkpoint) machinery from the kinetochore as they become properly attached to microtubules [14]. A recently identified protein, spindly, is needed for kinetochore recruitment of dynactin in some organisms, but not others (reviewed in [14]). ZW10 is part of a kinetochore-associated complex, RZZ (ROD/ZW10/Zwilch), which also recruits dynactin to the kinetochore and may work in combination with spindly [14,46]. However, ZW10 also interacts directly with a phosphorylated form of IC that is found only on unattached kinetochores, and this association is needed for dynein recruitment [61]. It has been proposed that, as kinetochores attach to the spindle and come under tension, IC is dephosphorylated, making it able to interact with p150 and triggering dynein-mediated removal of SAC components from the kinetochore [61].

LIS1, Nde1 and Ndel1 are also required for dynein function at the kinetochore. So far, it is not completely clear as to whether they act by recruiting dynein to kinetochores or by regulating dynein activity during microtubule attachment and SAC inactivation, or a combination of both. Again, the exact details vary between systems (reviewed in [14]). CENP-F (centromere protein F) targets Nde1 and Ndel1 to the kinetochore [62,63], independently of dynactin [62]. Nde1 and Ndel1 appear to have distinct functions at the kinetochore and during mitosis [63,64], with only Nde1 being needed for dynein recruitment [63]. Whether LIS1 is needed for Nde1/Ndel1 kinetochore association or vice versa, and how the microtubule plus end-binding protein CLIP170 (CAP-Gly domain-containing linker protein 170) is involved, is again system-dependent [14]. While RZZ is clearly needed for dynactin localization to kinetochores, disrupting its function has little or no effect on kinetochore Nde1 or Ndel1 levels or vice versa [62,65]. Dynein recruitment and function at the kinetochore is demonstrably complex, and further work is needed to determine the full network of interactions in each cell type or species.

Dynein association with membranous cargoes is similarly complicated. As outlined above and in Table 1, a range of membrane proteins have been identified that bind to dynein subunits. In addition, dynactin has also been thought to play a crucial role in dynein recruitment [34]. Attachment of dynactin to membranes has been proposed to be via the Arp1 (actin-related protein 1) filament binding to βIII spectrin (Table 2) [66,67]. In addition, several interactions between membrane proteins and the C-terminus of p150 have been discovered (Table 2). Interestingly, this same region of p150 is proposed to control the ability of dynactin to bind to membranes [68]. Furthermore, dynactin p50 binds to ZW10, which in interphase is part of a protein complex involved in trafficking between the ER (endoplasmic reticulum) and Golgi apparatus [14]. These results hint at a direct mechanistic link between dynactin, dynein and protein sorting in the secretory and endocytic pathways. Surprisingly, however, dynactin may actually play only a minor role in recruiting dynein, since dynein is still membrane-bound in Drosophila mutants lacking Arp1 [69]. Nevertheless, membrane movement in both directions is profoundly inhibited in Arp1 mutants [69], reinforcing the importance of dynactin for dynein function, and also underlining a close tie between dynein and kinesin activity (see below).

Table 2
Selected dynactin–cargo interactions

A range of key dynactin–cargo interactions is listed, along with the small GTPase involved, where appropriate. BPAG1n4, bullous pemphigoid antigen 1n4; ERGIC, endoplasmic reticulum–Golgi intermediate compartment; ORP1L, OSBP (oxysterol-binding protein)-related protein 1L; RILP, Rab-interacting lysosomal protein.

Dynactin subunit Cargo GTPase Other adaptor molecules Reference(s) 
p150 C-terminal COPII (coatamer protein II)-coated vesicles  Sec23p [121
p150 C-terminal Neuronal endosomes  BPAG1n4, via retrolinkin [122,123
p150 C-terminal Lysosomes Rab7 RILP, ORP1L, spectrin [66,124
p150 C-terminal Early endosomes  Sorting nexins 5 and 6 [125,126
p50 Kinetochores  ZW10 (RZZ) [14,46
p50 Secretory vesicles, ERGIC, Golgi Rab6 BicD1/2 [7779
Arp1 Golgi  βIII spectrin [67
Dynactin subunit Cargo GTPase Other adaptor molecules Reference(s) 
p150 C-terminal COPII (coatamer protein II)-coated vesicles  Sec23p [121
p150 C-terminal Neuronal endosomes  BPAG1n4, via retrolinkin [122,123
p150 C-terminal Lysosomes Rab7 RILP, ORP1L, spectrin [66,124
p150 C-terminal Early endosomes  Sorting nexins 5 and 6 [125,126
p50 Kinetochores  ZW10 (RZZ) [14,46
p50 Secretory vesicles, ERGIC, Golgi Rab6 BicD1/2 [7779
Arp1 Golgi  βIII spectrin [67

As is the case for kinetochores, LIS1, Nde1 and Ndel1 also play important roles in supporting dynein-driven positioning and movement of membranes [64,7074]. While this could be by regulating dynein's ATPase and force-generating properties, there is a simpler explanation: depletion of Nde1 and Ndel1 leads to a profound loss of dynein from membranes [64]. In contrast with the situation at the kinetochore, Nde1 and Ndel1 act redundantly in organelle positioning [64]. No membrane receptors for Nde1 and Ndel1 have been identified as yet, but they could conceivably associate directly with the lipid bilayer, since they are palmitoylated [75]. Knockdown of LIS1 also removed dynein, but to a lesser extent [64]. Interestingly, LIS1 is a subunit of the phospholipase PAFAH (platelet-activating factor acetylhydrolase) Ib, which is Golgi-localized and controls the formation of membrane tubules [71]. Loss of LIS1 could therefore affect Golgi apparatus morphology and position in two ways: via inhibition of dynein activity and its loss from the membrane, and by reduction in phospholipase activity. This could explain why LIS1 depletion leads to greater Golgi apparatus fragmentation than loss of Nde1 and Ndel1 [64]. Nde1 and Ndel1 appear to act upstream of LIS1, however, as LIS1 depletion had no effect on levels of Nde1 and Ndel1 on membranes [64].

The final dynein cargo adaptor I will describe is BicD. It was identified in Drosophila, and has since been shown to play a role in dynein-driven transport of mRNA, Rab6-positive membranes, lipid droplets and the nuclear envelope in higher eukaryotes [14]. It binds to dynein and dynactin p50, and to cargo-specific components such as Rab6, Egalitarian and the nuclear pore protein Ran-BP2 (Tables 1 and 2) [14,76]. In turn, Rab6 interacts with dynactin p150, dynein LC7 and membranes [14,77,78] while Egalitarian associates with mRNA and dynein LC8 [14]. However, the dynein-binding region of BicD when targeted artificially to mitochondria was sufficient to recruit dynein [79]. Interestingly, BicD2 also binds the kinesin-1 motor subunit [76,80] (see below).

Although the results described above paint a rather confusing picture of dynein–cargo interactions, the most straightforward interpretation is that dynein uses more than one mechanism to bind to most cargoes. These may include direct coupling between dynein and cargo molecules, and indirect associations brought about via adaptors such as dynactin or BicD, and the accessory factors Nde1 and Ndel1. A final example that underscores this complexity is dynein's binding to the nuclear envelope. This is required to maintain association between the nucleus and the MTOC, to establish correct nuclear position within the cell, especially during neuronal migration, and to facilitate nuclear envelope breakdown at the end of prophase (reviewed in [9,10]). SUN family members in the inner nuclear envelope provide a structural link from the nuclear lamina or chromosomes through to outer nuclear envelope proteins of the KASH family [9,10]. KASH proteins in turn can link directly to dynein, or indirectly via BicD or Nde1 (Table 1; [8183]). In an interesting parallel with kinetochores, CENP-F provides an alternative means of recruiting Nde1/Ndel1 (and hence dynein and LIS1) to the nuclear envelope via its interaction with the nuclear pore protein Nup133 [84]. The recruitment of dynein, LIS1 and Ndel1 to the nuclear envelope also depends on functional dynactin [85,86]. A third route for securing dynein to the nucleus is provided by BicD2, which is recruited to the nuclear envelope by binding to the nuclear pore component Ran-BP2 [76]. A final twist is that KASH proteins, Ran-BP2 and BicD all bind to kinesin-1, which is needed to enable bidirectional movement of the nucleus and the MTOC [9,10,76,82,87]. As discussed in the following section, bidirectional transport is common within the cell, generating important questions about how dynein function is controlled.

Regulation of dynein function in the cell

Since motor proteins in the cell fulfil such a wide range of functions in so many different contexts, they must clearly be regulated in some way. For example, any given cargo may be moving or not, and this control could be exerted over time (such as through the cell cycle) or in concert with other cellular functions, such as a particular membrane trafficking step. Indeed, as mentioned above, there are links between dynein, dynactin and important trafficking components (Tables 1 and 2). It is likely, too, that dynein will be regulated independently on different cargoes.

Two obvious possibilities exist for modulating dynein-dependent movement, and there is evidence for both. One is that active dynein is recruited when it is needed, then removed again when movement is complete. The second is that dynein is bound at all times, but can be switched between active and inactive states.

Since dynein generally transports cargo towards the cell centre, and is also active at the cell cortex, it would be useful to have a means of enriching dynein in the cell periphery ready for action. The accumulation of dynein and dynactin on the plus ends of growing microtubules achieves this in many organisms [14]. In yeast, delivery of dynein to the bud cortex via attachment to growing microtubules is important for subsequent transport of the nucleus into the bud during mitosis [14]. In filamentous fungi, dynein is transported to microtubule plus ends by kinesin-1 [88,89] where interactions with dynactin help it accumulate [90]. Early endosomes that move towards plus ends using kinesin-3 were shown to lack dynein [28], and it was initially proposed that they picked up dynein from the plus end pool, ready for the return journey [89]. However, more recent work has shown that plus-end-directed endosomes often switched direction before reaching the tip by recruiting dynein molecules that were moving towards them along the same microtubule [28].

In vertebrates, dynein is rarely seen at microtubule plus ends, while dynactin clearly behaves as a +TIP (microtubule plus-end-interacting protein) [14,91]. It has been suggested that contact between a growing microtubule and a stationary membrane cargo delivers dynactin, thus initiating dynein-driven movement [91]. However, other work has shown that loss of p150 from plus ends has no effect on membrane movement in vertebrates [92,93]. Instead, the +TIP CLIP170 was seen to activate motility of Xenopus melanosomes [92], but not of secretory and endocytic organelles [93].

Analysis of cargoes that move bidirectionally has provided support for the second model, where activity is regulated rather than cargo association. For example, GFP (green fluorescent protein)-tagged dynein localizes to particles that move in both directions along microtubules in living cells (e.g. [42,52,94,95]). Furthermore, prion-protein-containing vesicles possess similar levels of dynein and kinesin-1 in vivo, irrespective of which direction they are moving, or indeed whether they are moving at all [96], while vesicles labelled with GFP–p50/dynamitin move bidirectionally in vitro [27].

How bidirectional cargoes switch their direction of movement has fascinated many researchers (reviewed in [97]). One proposal is that both motors are always active and participate in a tug-of-war [26,27,97]. In vertebrates and Dictyostelium, several relatively weak dyneins compete against single stronger kinesins [26,27], whereas in Ustilago maydis, one dynein competes against four to five kinesin-3 molecules [28]. However, a tug-of-war seems rather a wasteful mechanism, and a refinement of this model is that the competition only lasts until one motor wins, and, after that, the loser is inactivated [97,98]. Alternatively, opposing forces could be harnessed to deform membranes such as endosomes in a way that facilitates sorting events [26]. The number of each motor per cargo will clearly be important in determining the bias in direction of movement, and BicD and Egalitarian levels have been shown to be influence the amount of dynein loaded on to mRNPs in Drosophila [99].

In the simplest tug-of-war model, if one motor is turned off, the other motor will always win. However, in many instances this is not the case; inhibition of one direction of movement leads to inactivation of the other direction as well [40,69,97,98]. The clearest demonstration of this is peroxisome movement, where loss of kinesin-1 activity inhibits dynein-driven translocations as well [40,98]. However, replacement of kinesin-1 with any functional plus-end motor that is artificially targeted to the peroxisome will reactivate dynein, even if the artificial motor moves far more slowly than kinesin-1 [98]. This suggests that the generation of force in the plus-end direction is all that is needed to activate dynein, perhaps by a tension-sensing mechanism [98,100].

How might this tension-based coupling be achieved? One possibility is that the presence of two opposing motors within a small structure is sufficient, as force generated by one will necessarily exert tension on the other. This idea is supported by the apparently contradictory finding that kinesin-1 and dynein act independently of each other in the ER [101]; the ER is such a large, dynamic and deformable membrane structure that dynein and kinesin may simply end up in distinct regions of the organelle, acting independently.

An alternative means of communicating tension between motors is for them to be physically associated. This may well be the case, at least in some circumstances, since IC has been shown to bind to kinesin LC [102]. Indirect associations are also likely via scaffolding complexes. For example, dynactin also interacts with kinesin-2 and kinesin-5 [14], but whether it plays a role in co-ordinating motor switching is not clear. Furthermore, both Ran-BP2 and BicD interact with kinesin-1 heavy chain [76,80] while BicD-related protein 1 binds Kif1C [103], and BicD levels have been shown to affect both directions of lipid droplet movement [104]. In addition, there is considerable evidence that JIPs (c-Jun N-terminal kinase-interacting proteins), originally identified as kinesin-1 binding proteins, are involved in regulating dynein-driven movement in neurons (Table 1; [105107]) and non-neuronal cells [108].

While a tug-of-war may control the direction of movement over short time-scales, there is clearly scope for longer-term changes that involve motor recruitment/loss or activation/inactivation [27,98]. How dynactin, LIS1, Nde1, Ndel1 and BicD contribute to this regulation remains to be determined in detail. Additional regulatory molecules such as huntingtin [109], and a variety of GTPases (Tables 1 and 2; [14,110,111]) are also likely to influence dynein activity. Phosphorylation is another obvious means of controlling dynein function, but again, detailed mechanisms are lacking.

Conclusion

Recent years have seen great progress in understanding dynein's structure and mechanism, and the contribution made by accessory proteins to its function. We now know much more about how dynein is targeted to cargoes, but this has added further complexity, given that there are often several parallel sets of molecules that contribute. Our knowledge of how dynein is regulated in vivo is still fairly limited, however, and it seems likely that systematic studies which look at the whole range of potential regulators together will be needed to unravel the details of control mechanisms.

Cellular Cytoskeletal Motor Proteins: A Biochemical Society/Wellcome Trust Focused Meeting held at Wellcome Trust Genome Campus, Hinxton, Cambridge, U.K., 30 March–1 April 2011. Organized and Edited by Folma Buss (Cambridge, U.K.) and John Kendrick-Jones (MRC Laboratory of Molecular Biology, Cambridge, U.K.).

Abbreviations

     
  • AAA+

    ATPase associated with various cellular activities

  •  
  • Arp1

    actin-related protein 1

  •  
  • BicD

    Bicaudal D

  •  
  • CAP-Gly

    cytoskeleton-associated protein glycine-rich

  •  
  • CENP-F

    centromere protein F

  •  
  • CLIP170

    CAP-Gly domain-containing linker protein 170

  •  
  • DHC

    dynein heavy chain

  •  
  • ER

    endoplasmic reticulum

  •  
  • GFP

    green fluorescent protein

  •  
  • IC

    intermediate chain

  •  
  • JIP

    c-Jun N-terminal kinase-interacting proteins

  •  
  • LC

    light chain

  •  
  • LIC

    light intermediate chain

  •  
  • LIS1

    lissencephaly 1

  •  
  • mRNP

    messenger ribonucleoprotein

  •  
  • MTOC

    microtubule-organizing centre

  •  
  • NudE

    nuclear distribution protein E

  •  
  • RZZ

    ROD/ZW10/Zwilch

  •  
  • SAC

    spindle assembly checkpoint

  •  
  • +TIP

    microtubule plus-end-interacting protein

  •  
  • ZW10

    Zest White 10

I thank Philip Woodman and Alan Roseman for helpful discussion and comments on the paper. I apologize to the many people whose work I have been unable to cite due to space constraints.

Funding

This work was funded by The Wellcome Trust [grant number 086077/Z/08/Z], The Medical Research Council [grant number G0900930] and the Biotechnology and Biological Sciences Research Council [grant number BB/H017828/1].

References

References
1
Pfister
K.K.
Shah
P.
Hummerich
H.
Russ
A.
Cotton
J.
Annuar
A.
King
S.J.
Fisher
E.M.
Genetic analysis of the cytoplasmic dynein subunit families
PLoS Genet.
2006
, vol. 
2
 pg. 
e1
 
2
Palmer
K.J.
Hughes
H.
Stephens
D.J.
Specificity of cytoplasmic dynein subunits in discrete membrane trafficking steps
Mol. Biol. Cell
2009
, vol. 
20
 (pg. 
2885
-
2899
)
3
Wickstead
B.
Gull
K.
Dyneins across eukaryotes: a comparative genomic analysis
Traffic
2007
, vol. 
8
 (pg. 
1708
-
1721
)
4
Zheng
Y.
Wildonger
J.
Ye
B.
Zhang
Y.
Kita
A.
Younger
S.
Zimmerman
S.
Jan
L.
Jan
Y.
Dynein is required for polarized dendritic transport and uniform microtubule orientation in axons
Nat. Cell Biol.
2008
, vol. 
10
 (pg. 
1172
-
1180
)
5
Satoh
D.
Sato
D.
Tsuyama
T.
Saito
M.
Ohkura
H.
Rolls
M.
Ishikawa
F.
Uemura
T.
Spatial control of branching within dendritic arbors by dynein-dependent transport of Rab5-endosomes
Nat. Cell Biol.
2008
, vol. 
10
 (pg. 
1164
-
1171
)
6
Bianco
A.
Dienstbier
M.
Salter
H.
Gatto
G.
Bullock
S.
Bicaudal-D regulates fragile X mental retardation protein levels, motility and function during neuronal morphogenesis
Curr. Biol.
2010
, vol. 
20
 (pg. 
1487
-
1492
)
7
Kapitein
L.C.
Schlager
M.
Kuijpers
M.
Wulf
P.
van Spronsen
M.
MacKintosh
F.
Hoogenraad
C.C.
Mixed microtubules steer dynein-driven cargo transport into dendrites
Curr. Biol.
2010
, vol. 
20
 (pg. 
290
-
299
)
8
Hirokawa
N.
Niwa
S.
Tanaka
Y.
Molecular motors in neurons: transport mechanisms and roles in brain function, development and disease
Neuron
2010
, vol. 
68
 (pg. 
610
-
638
)
9
Starr
D.
A nuclear-envelope bridge positions nuclei and moves chromosomes
J. Cell Sci.
2009
, vol. 
122
 (pg. 
577
-
586
)
10
Burke
B.
Roux
K.
Nuclei take a position: managing nuclear location
Dev. Cell
2009
, vol. 
17
 (pg. 
587
-
597
)
11
Wynshaw-Boris
A.
Pramparo
T.
Youn
Y.
Hirotsune
S.
Lissencephaly: mechanistic insights from animal models and potential therapeutic strategies
Semin. Cell Dev. Biol.
2010
, vol. 
21
 (pg. 
823
-
830
)
12
Hafezparast
M.
Klocke
R.
Ruhrberg
C.
Murquardt
A.
Ahmad-Annuar
A.
Bowen
S.
Lalli
G.
Witherden
A.
Hummerich
H.
Nicholson
S.
, et al. 
Mutations in dynein link motor neuron degeneration to defects in retrograde transport
Science
2003
, vol. 
300
 (pg. 
808
-
812
)
13
Banks
G.
Haas
M.
Line
S.
Shepherd
H.
AlQatari
M.
Stewart
S.
Rishal
I.
Philpott
A.
Kalmar
B.
Kuta
A.
, et al. 
Behavioral and other phenotypes in a cytoplasmic dynein light intermediate chain 1 mutant mouse
J. Neurosci.
2011
, vol. 
31
 (pg. 
5483
-
5494
)
14
Kardon
J.
Vale
R.
Regulators of the cytoplasmic dynein motor
Nat. Rev. Mol. Cell Biol.
2009
, vol. 
10
 (pg. 
854
-
865
)
15
Gee
M.
Heuser
J.E.
Vallee
R.
An extended microtubule-binding structure within the dynein motor domain
Nature
1997
, vol. 
390
 (pg. 
636
-
639
)
16
Roberts
A.
Numata
A.
Walker
M.
Kato
Y.
Malkova
B.
Kon
T.
Ohkura
R.
Arisaka
F.
Knight
P.J.
Sutoh
K.
Burgess
S.
AAA+ ring and linker swing mechanism in the dynein motor
Cell
2009
, vol. 
136
 (pg. 
485
-
495
)
17
Kon
T.
Sutoh
K.
Kurisu
G.
X-ray structure of a functional full-length dynein motor domain
Nat. Struct. Mol. Biol.
2011
, vol. 
18
 (pg. 
638
-
642
)
18
Carter
A.P.
Cho
C.
Jin
L.
Vale
R.
Crystal structure of the dynein motor domain
Science
2011
, vol. 
331
 (pg. 
1159
-
1165
)
19
Houdusse
A.
Carter
A.P.
Dynein swings into action
Cell
2009
, vol. 
136
 (pg. 
395
-
396
)
20
Ori-McKenney
K.
Xu
J.
Gross
S.
Vallee
R.
A cytoplasmic dynein tail mutation impairs motor processivity
Nat. Cell Biol.
2010
, vol. 
12
 (pg. 
1228
-
1234
)
21
Nan
X.
Sims
P.A.
Chen
P.
Xie
X.S.
Observation of individual microtubule motor steps in living cells with endocytosed quantum dots
J. Phys. Chem. B
2005
, vol. 
109
 (pg. 
24220
-
24224
)
22
Gennerich
A.
Carter
A.P.
Reck-Peterson
S.L.
Vale
R.D.
Force-induced bidirectional stepping of cytoplasmic dynein
Cell
2007
, vol. 
131
 (pg. 
952
-
965
)
23
Mallik
R.
Carter
B.C.
Lex
S.A.
King
S.J.
Gross
S.P.
Cytoplasmic dynein functions as a gear in response to load
Nature
2004
, vol. 
427
 (pg. 
649
-
652
)
24
King
S.
Schroer
T.
Dynactin increases the processivity of the cytoplasmic dynein motor
Nat. Cell Biol.
1999
, vol. 
2
 (pg. 
20
-
24
)
25
Kural
C.
Kim
H.
Syed
S.
Goshima
G.
Gelfand
V.
Selvin
P.
Kinesin and dynein move a peroxisome in vivo: a tug-of-war or coordinated movement?
Science
2005
, vol. 
308
 (pg. 
1469
-
1472
)
26
Soppina
V.
Rai
A.
Ramaiya
A.
Barak
P.
Mallik
R.
Tug-of-war between dissimilar teams of microtubule motors regulates transport and fission of endosomes
Proc. Natl. Acad. Sci. U.S.A.
2009
, vol. 
106
 (pg. 
19381
-
19386
)
27
Hendricks
A.
Perlson
E.
Ross
J.
Schroeder
H.
III
Tokito
M.
Holzbaur
E.L.F.
Motor coordination via a tug-of-war mechanism drives bidirectional vesicle transport
Curr. Biol.
2010
, vol. 
20
 (pg. 
697
-
702
)
28
Schuster
M.
Lipowsky
R.
Assmann
M.-A.
Lenz
P.
Steinberg
G.
Transient binding of dynein controls bidirectional long-range motility of early endosomes
Proc. Natl. Acad. Sci. U.S.A.
2011
, vol. 
108
 (pg. 
3618
-
3623
)
29
Shubeita
G.
Tran
S.
Xu
J.
Vershinin
M.
Cermelli
S.
Cotton
S.
Welte
M.A.
Gross
S.P.
Consequences of motor copy number on the intracellular transport of kinesin-1-driven lipid droplets
Cell
2008
, vol. 
135
 (pg. 
1098
-
1107
)
30
Martinez
J.E.
Vershinin
M.D.
Shubeita
G.T.
Gross
S.P.
On the use of in vivo cargo velocity as a biophysical marker
Biochem. Biophys. Res. Commun.
2007
, vol. 
353
 (pg. 
835
-
840
)
31
Ross
J.
Shuman
H.
Holzbaur
E.
Goldman
Y.
Kinesin and dynein-dynactin at intersecting microtubules: motor density affects dynein function
Biophys. J.
2008
, vol. 
94
 (pg. 
3115
-
3125
)
32
Reck-Peterson
S.
Yildiz
A.
Carter
A.
Gennerich
A.
Zhang
N.
Vale
R.
Single-molecule analysis of dynein processivity and stepping behavior
Cell
2006
, vol. 
126
 (pg. 
335
-
348
)
33
Ross
J.
Wallace
K.
Shuman
H.
Goldman
Y.
Holzbaur
E.
Processive bidirectional motion of dynein–dynactin complexes in vitro
Nat. Cell Biol.
2006
, vol. 
8
 (pg. 
562
-
570
)
34
Schroer
T.
Dynactin
Annu. Rev. Cell Dev. Biol.
2004
, vol. 
20
 (pg. 
759
-
779
)
35
Culver-Hanlon
T.
Lex
S.
Stephens
A.
Quintyne
N.
King
S.
A microtubule-binding domain in dynactin increases dynein processivity by skating along microtubules
Nat. Cell Biol.
2006
, vol. 
8
 (pg. 
264
-
270
)
36
Gross
S.
Tuma
M.
Deacon
S.
Serpinskaya
A.
Reilein
A.
Gelfand
V.
Interactions and regulation of molecular motors in Xenopus melanophores
J. Cell Biol.
2002
, vol. 
156
 (pg. 
855
-
865
)
37
Kobayashi
T.
Shiroguchi
K.
Edamatsu
M.
Toyoshima
Y.
Microtubule-binding properties of p150 expedient for dynein motility
Biochem. Biophys. Res. Commun.
2006
, vol. 
340
 (pg. 
23
-
28
)
38
Waterman-Storer
C.M.
Karki
S.
Holzbaur
E.L.
The p150Glued component of the dynactin complex binds to both microtubules and the actin-related protein centractin (Arp-1)
Proc. Natl. Acad. Sci. U.S.A.
1995
, vol. 
92
 (pg. 
1634
-
1638
)
39
Dixit
R.
Levy
J.
Tokito
M.
Ligon
L.
Holzbaur
E.
Regulation of dynactin through the differential expression of p150Glued isoforms
J. Biol. Chem.
2008
, vol. 
283
 (pg. 
33611
-
33619
)
40
Kim
H.
Ling
S.-C.
Rogers
G.
Kural
C.
Selvin
P.
Rogers
S.
Gelfand
V.
Microtubule binding by dynactin is required for microtubule organization but not cargo transport
J. Cell Biol.
2007
, vol. 
176
 (pg. 
641
-
651
)
41
Kardon
J.
Reck-Peterson
S.
Vale
R.
Regulation of the processivity and intracellular localization of Saccharomyces cerevisiae dynein by dynactin
Proc. Natl. Acad. Sci. U.S.A.
2009
, vol. 
106
 (pg. 
5669
-
5674
)
42
Yamada
M.
Toba
S.
Yoshida
Y.
Haratani
K.
Mori
D.
Yano
Y.
Mimori-Kiyosue
Y.
Nakamura
T.
Itoh
K.
Fushiki
S.
, et al. 
LIS1 and NDEL1 coordinate the plus-end-directed transport of cytoplasmic dynein
EMBO J.
2008
, vol. 
27
 (pg. 
2471
-
2483
)
43
McKenney
R.
Vershinin
M.
Kunwar
A.
Vallee
R.
Gross
S.
LIS1 and NudE induce a persistent dynein force-producing state
Cell
2010
, vol. 
141
 (pg. 
304
-
314
)
44
Tai
C.-Y.
Dujardin
D.
Faulkner
N.
Vallee
R.
Role of dynein, dynactin and CLIP-170 interactions in LIS1 kinetochore function
J. Cell Biol.
2002
, vol. 
156
 (pg. 
959
-
968
)
45
Sasaki
S.
Shionoya
A.
Ishida
M.
Gambello
M.
Yingling
J.
Wynshaw-Boris
A.
Hirotsune
S.
A LIS1/NUDEL/cytoplasmic dynein heavy chain complex in the developing and adult nervous system
Neuron
2000
, vol. 
28
 (pg. 
681
-
696
)
46
Stehman
S.
Chen
Y.
McKenny
R.
Vallee
R.
NudE and NudEL are required for mitotic progression and are involved in dynein recruitment to kinetochores
J. Cell Biol.
2007
, vol. 
178
 (pg. 
583
-
594
)
47
Zylkiewicz
E.
Kijanska
M.
Choi
W.-C.
Derewenda
U.
Derewenda
Z.
Stukenberg
P.T.
The N-terminal coiled-coil of Ndel1 is a regulated scaffold that recruits LIS1 to dynein
J. Cell Biol.
2011
, vol. 
192
 (pg. 
433
-
445
)
48
Wang
S.
Zheng
Y.
Identification of a novel dynein binding domain in Nudel essential for spindle pole organization in Xenopus egg extract
J. Biol. Chem
2011
, vol. 
286
 (pg. 
587
-
593
)
49
Torisawa
T.
Nakayama
A.
Furuta
K.
Yamada
M.
Hirotsune
S.
Toyoshima
Y.
Functional dissection of LIS1 and NDEL1 towards understanding the molecular mechanisms of cytoplasmic dynein regulation
J. Biol. Chem.
2011
, vol. 
286
 (pg. 
1959
-
1965
)
50
Vaughan
P.
Lesyzk
J.
Vaughan
K.
Cytoplasmic dynein intermediate chain phosphorylation regulates binding to dynactin
J. Biol. Chem.
2001
, vol. 
276
 (pg. 
26171
-
26179
)
51
Kuta
S.
Deng
W.
El-Kadi
A.
Banks
G.
Hafezparast
M.
Pfister
K.
Fisher
E.
Mouse cytoplasmic dynein intermediate chains: identification of new isoforms, alternative splicing and tissue distribution of transcripts
PLoS ONE
2010
, vol. 
5
 pg. 
e11682
 
52
Ha
J.
Lo
K.-H.
Myers
K.
Carr
T.
Humsi
M.
Rasoul
B.
Segal
R.A.
Pfister
K.
A neuron-specific cytoplasmic dynein isoform preferentially transports TrkB signaling endosomes
J. Cell Biol.
2008
, vol. 
181
 (pg. 
1027
-
1039
)
53
Tynan
S.H.
Purohit
A.
Doxsey
S.J.
Vallee
R.B.
Light intermediate chain 1 defines a functional subfraction of cytoplasmic dynein which binds to pericentrin
J. Biol. Chem.
2000
, vol. 
275
 (pg. 
32763
-
32768
)
54
Schmoranzer
J.
Fawcett
J.P.
Segura
M.
Tan
S.
Vallee
R.B.
Pawson
T.
Gundersen
G.G.
Par3 and dynein associate to regulate local microtubule dynamics and centrosome orientation during migration
Curr. Biol.
2009
, vol. 
19
 (pg. 
1065
-
1074
)
55
Reilen
A.
Serpinskaya
A.
Karcher
R.
Dujardin
D.
Vallee
R.
Gelfand
V.
Differential regulation of dynein-driven melanosome movement
Biochem. Biophys. Res. Commun.
2003
, vol. 
309
 (pg. 
652
-
658
)
56
Tan
S.
Scherer
J.
Vallee
R.
Recruitment of dynein to late endosomes and lysosomes through light intermediate chains
Mol. Biol. Cell
2011
, vol. 
22
 (pg. 
467
-
477
)
57
Horgan
C.P.
Hanscom
S.R.
Jolly
R.S.
Futter
C.E.
McCaffrey
M.W.
Rab11–FIP3 links the Rab11 GTPase and cytoplasmic dynein to mediate transport to the endosomal-recycling compartment
J. Cell Sci.
2010
, vol. 
123
 (pg. 
181
-
191
)
58
Horgan
C.P.
Hanscom
S.R.
Jolly
R.S.
Futter
C.E.
McCaffrey
M.W.
Rab11–FIP3 binds dynein light intermediate chain 2 and its overexpression fragments the Golgi complex
Biochem. Biophys. Res. Commun.
2010
, vol. 
394
 (pg. 
387
-
392
)
59
Sivaram
M.V.
Wadzinski
T.L.
Redick
S.D.
Manna
T.
Doxsey
S.J.
Dynein light intermediate chain 1 is required for progress through the spindle assembly checkpoint
EMBO J.
2009
, vol. 
28
 (pg. 
902
-
914
)
60
Echeverri
C.J.
Paschal
B.M.
Vaughan
K.T.
Vallee
R.B.
Molecular characterisation of the 50-kD subunit of dynactin reveals function for the complex in chromosome alignment and spindle organisation during mitosis
J. Cell Biol.
1996
, vol. 
132
 (pg. 
617
-
633
)
61
Whyte
J.
Bader
J.
Tauhata
S.
Raycroft
M.
Hornick
J.
Pfister
K.
Lane
W.
Chan
G.K.
Hincliffe
E.
Vaughan
P.S.
Vaughan
K.T.
Phosphorylation regulates targeting of cytoplasmic dynein to kinetochores during mitosis
J. Cell Biol.
2008
, vol. 
183
 (pg. 
819
-
834
)
62
Liang
Y.
Yu
W.
Li
Y.
Yu
L.
Zhang
Q.
Wang
F.
Yang
Z.
Du
J.
Huang
Q.
Yao
X.
Zhu
X.
Nudel modulates kinetochore association and function of cytoplasmic dynein in M phase
Mol. Biol. Cell
2007
, vol. 
18
 (pg. 
2656
-
2666
)
63
Vergnolle
M.
Taylor
S.
Cenp-F links kinetochores to Ndel1/Nde1/Lis1/dynein microtubule motor complexes
Curr. Biol.
2007
, vol. 
17
 (pg. 
1173
-
1179
)
64
Lam
C.
Vergnolle
M.A.
Thorpe
L.
Woodman
P.
Functional interplay between LIS1, NDE1 and NDEL1 in dynein-dependent organelle positioning
J. Cell Sci.
2010
, vol. 
123
 (pg. 
202
-
212
)
65
Wainman
A.
Creque
J.
Williams
B.C.
Williams
E.
Bonaccorsi
S.
Gatti
M.
Goldberg
M.L.
Roles of Drosophila NudE protein in kinetochore function and centrosome migration
J. Cell Sci.
2009
, vol. 
122
 (pg. 
1747
-
1758
)
66
Johansson
M.
Rocha
N.
Zwart
W.
Jordens
I.
Janssen
L.
Kuijl
C.
Olkkonen
V.
Neefjes
J.
Activation of endosomal dynein motors by stepwise assembly of Rab7–RILP–p150Glued, ORP1L and the receptor βIII spectrin
J. Cell Biol.
2007
, vol. 
176
 (pg. 
459
-
471
)
67
Holleran
E.A.
Tokito
M.K.
Karki
S.
Holzbaur
E.L.F.
Centractin (ARP1) associates with spectrin revealing a potential mechanism to link dynactin to intracellular organelles
J. Cell Biol.
1996
, vol. 
135
 (pg. 
1815
-
1829
)
68
Kumar
S.
Zhou
Y.
Plamann
M.
Dynactin–membrane interaction is regulated by the C-terminal domains of p150Glued
EMBO Rep.
2001
, vol. 
2
 (pg. 
939
-
944
)
69
Haghnia
M.
Cavalli
V.
Shah
S.
Schimmelpfeng
K.
Brusch
R.
Herrera
C.
Pilling
A.
Goldstein
L.
Dynactin is required for coordinated bidirectional motility, but not for dynein membrane attachment
Mol. Biol. Cell
2007
, vol. 
18
 (pg. 
2081
-
2089
)
70
Liang
Y.
Yu
W.
Li
Y.
Yang
Z.
Yan
X.
Huang
Q.
Zhu
X.
Nudel functions in membrane traffic mainly through association with Lis1 and cytoplasmic dynein
J. Cell Biol.
2004
, vol. 
164
 (pg. 
557
-
566
)
71
Bechler
M.
Doody
A.
Racoosin
E.
Lin
L.
Lee
K.
Brown
W.J.
The phospholipase complex PAFAH Ib regulates the functional organization of the Golgi complex
J. Cell Biol.
2010
, vol. 
190
 (pg. 
45
-
53
)
72
Ding
C.
Liang
X.
Ma
L.
Yuan
X.
Zhu
X.
Opposing effects of Ndel1 and α1 or α2 on cytoplasmic dynein through competitive binding to Lis1
J. Cell Sci.
2009
, vol. 
122
 (pg. 
2820
-
2827
)
73
Sasaki
S.
Mori
D.
Toyo-oka
K.
Chen
A.
Garrett-Beal
L.
Muramutsu
M.
Miyagawa
S.
Hiraiwa
N.
Yoshiki
A.
Wynshaw-Boris
A.
Hirotsune
S.
Complete loss of Ndel1 results in neuronal migration defects and early embryonic lethality
Mol. Cell. Biol.
2005
, vol. 
25
 (pg. 
7812
-
7827
)
74
Guo
J.
Yang
Z.
Song
W.
Chen
Q.
Wang
F.
Zhang
Q.
Zhu
X.
Nudel contributes to microtubule anchoring at the mother centriole and is involved in both dynein-dependent and -independent centrosomal protein assembly
Mol. Biol. Cell
2006
, vol. 
17
 (pg. 
680
-
689
)
75
Shmueli
A.
Segal
M.
Sapir
T.
Tsutsumi
R.
Noritake
J.
Bar
A.
Sapoznik
S.
Fukata
Y.
Orr
I.
Fukata
M.
Reiner
O.
Ndel1 palmitoylation: a new mean to regulate cytoplasmic dynein activity
EMBO J.
2010
, vol. 
29
 (pg. 
107
-
119
)
76
Splinter
D.
Tanenbaum
M.
Lindqvist
A.
Jaarsman
D.
Flotho
A.
Yu
K.
Grigoriev
I.
Engelsma
D.
Haasdijk
E.
Keijzer
N.
, et al. 
Bicaudal D2, dynein, and kinesin-1 associate with nuclear pore complexes and regulate centrosome and nuclear positioning during mitotic entry
PLoS Biol.
2010
, vol. 
8
 pg. 
e1000350
 
77
Matanis
T.
Akhmanova
A.
Wulf
P.
Del Nery
E.
Weide
T.
Stepanova
T.
Galjart
N.
Grosveld
F.
Goud
B.
de Zeeuw
C.
, et al. 
Bicaudal-D regulates COPI-independent Golgi–ER transport by recruiting the dynein–dynactin motor complex
Nat. Cell Biol.
2002
, vol. 
4
 (pg. 
986
-
992
)
78
Short
B.
Preisinger
C.
Schaletzky
J.
Kopajtich
R.
Barr
F.
The Rab6 GTPase regulates recruitment of the dynactin complex to Golgi membranes
Curr. Biol.
2002
, vol. 
12
 (pg. 
1792
-
1795
)
79
Hoogenraad
C.
Wulf
P.
Schiefermeier
N.
Stepanova
T.
Galjart
N.
Small
J.
Grosveld
F.
de Zeeuw
C.
Akhmanova
A.
Bicaudal D induces selective dynein-mediated microtubule minus end-directed transport
EMBO J.
2003
, vol. 
22
 (pg. 
6004
-
6015
)
80
Grigoriev
I.
Splinter
D.
Keijzer
N.
Wulf
P.
Demmers
J.
Ohtsuka
T.
Modesti
M.
Maly
I.
Grosveld
F.
Hoogenraad
C.C.
Akhmanova
A.
Rab6 regulates transport and targeting of exocytic carriers
Dev. Cell
2007
, vol. 
13
 (pg. 
305
-
314
)
81
Zhang
X.
Lei
K.
Yuan
X.
Wu
X.
Zhuang
Y.
Xu
T.
Xu
R.
Han
M.
SUN1/2 and Syne/Nesprin-1/2 complexes connect centrosome to the nucleus during neurogenesis and neuronal migration in mice
Neuron
2009
, vol. 
64
 (pg. 
173
-
187
)
82
Fridolfsson
H.
Ly
N.
Meyerzon
M.
Starr
D.
UNC-83 coordinates kinesin-1 and dynein activities at the nuclear envelope during nuclear migration
Dev. Biol.
2010
, vol. 
338
 (pg. 
237
-
250
)
83
Malone
C.
Misner
L.
Le Bot
N.
Tsai
M.-C.
Campbell
J.
Ahringher
J.
White
J.
The C. elegans hook protein, ZYG12, mediates the essential attachment between the centrosome and nucleus
Cell
2003
, vol. 
115
 (pg. 
825
-
836
)
84
Bolhy
S.
Bouhlel
I.
Dultz
E.
Nayak
T.
Zuccolo
M.
Gatti
X.
Vallee
R.
Ellenberg
J.
Doye
V.
A Nup133-dependent NPC-anchored network tethers centrosomes to the nuclear envelope in prophase
J. Cell Biol.
2011
, vol. 
195
 (pg. 
855
-
871
)
85
Hebbar
S.
Mesgnon
M.
Guillotte
A.
Desai
B.
Ayala
R.
Smith
D.
Lis1 and Ndel1 influence the timing of nuclear envelope breakdown in neural stem cells
J. Cell Biol
2008
, vol. 
182
 (pg. 
1063
-
1071
)
86
Salina
D.
Bodoor
K.
Eckley
D.
Schroer
T.A.
Rattner
J.B.
Burke
B.
Cytoplasmic dynein as a facilitator of nuclear envelope breakdown
Cell
2002
, vol. 
108
 (pg. 
97
-
107
)
87
Fridolfsson
H.
Starr
D.
Kinesin-1 and dynein at the nuclear envelope mediate the bidrectional migrations of nuclei
J. Cell Biol.
2010
, vol. 
191
 (pg. 
115
-
128
)
88
Zhang
J.
Li
S.
Fischer
R.
Xiang
X.
Accumulation of cytoplasmic dynein and dynactin at microtubule plus ends in Aspergillus nidulans is kinesin dependent
Mol. Biol. Cell
2003
, vol. 
14
 (pg. 
1479
-
1488
)
89
Lenz
J.
Schuchardt
I.
Straube
A.
Steinberg
G.
A dynein loading zone for retrograde endosome motility at microtubule plus ends
EMBO J.
2006
, vol. 
25
 (pg. 
2275
-
2286
)
90
Schuster
M.
Kilaru
S.
Ashwin
P.
Lin
C.
Severs
N.
Steinberg
G.
Controlled and stochastic retention concentrates dynein at microtubule ends to keep endosomes on track
EMBO J.
2011
, vol. 
30
 (pg. 
652
-
664
)
91
Vaughan
P.
Miura
P.
Henderson
M.
Byrne
B.
Vaughan
K.
A role for regulated binding of p150Glued to microtubule plus ends in organelle transport
J. Cell Biol.
2002
, vol. 
158
 (pg. 
305
-
319
)
92
Lomakin
A.
Semenova
I.V.
Zaliapin
I.
Kraikivski
P.
Nadezhdina
E.
Slepchenko
B.
Akhmanova
A.
Rodionov
V.
CLIP-170-dependent capture of membrane organelles by microtubules initiates minus-end directed transport
Dev. Cell
2009
, vol. 
17
 (pg. 
323
-
333
)
93
Watson
P.
Stephens
D.
Microtubule plus-end loading of p150Glued is mediated by EB1 and CLIP-170 but is not required for intracellular membrane traffic in mammalian cells
J. Cell Sci.
2006
, vol. 
119
 (pg. 
2758
-
2767
)
94
Ma
S.
Chisholm
R.
Cytoplasmic dynein-associated structures move bidirectionally in vivo
J. Cell Sci.
2002
, vol. 
115
 (pg. 
1453
-
1460
)
95
Kobayashi
T.
Marayama
T.
Cell cycle-dependent microtubule-based dynamic transport of cytoplasmic dynein in mammalian cells
PLoS ONE
2009
, vol. 
4
 pg. 
e7827
 
96
Encalada
S.
Szpankowksi
L.
Xia
C.-H.
Goldstein
L.
Stable kinesin and dynein assemblies drive the axonal transport of mammalian prion protein vesicles
Cell
2011
, vol. 
144
 (pg. 
551
-
565
)
97
Welte
M.
Bidirectional transport along microtubules
Curr. Biol.
2004
, vol. 
14
 (pg. 
R525
-
R537
)
98
Ally
S.
Larson
A.
Barlan
K.
Rice
S.
Gelfand
V.
Opposite-polarity motors activate one another to trigger cargo transport in live cells
J. Cell Biol.
2009
, vol. 
187
 (pg. 
1071
-
1082
)
99
Bullock
S.
Nicol
A.
Gross
S.
Zicha
D.
Guidance of bidirectional motor complexes by mRNA cargoes through control of dynein number and activity
Curr. Biol.
2006
, vol. 
16
 (pg. 
1447
-
1452
)
100
Howard
J.
Mechanical signaling in networks of motor and cytoskeletal proteins
Annu. Rev. Biophys.
2009
, vol. 
38
 (pg. 
217
-
234
)
101
Wozniak
M.
Bola
B.
Brownhill
K.
Yang
Y.-C.
Levakova
V.
Allan
V.
Role of kinesin-1 and cytoplasmic dynein in endoplasmic reticulum movement in VERO cells
J. Cell Sci.
2009
, vol. 
122
 (pg. 
1979
-
1989
)
102
Ligon
L.
Tokito
M.
Finkelstein
J.
Grossman
F.
Holzbaur
E.
A direct interaction between cytoplasmic dynein and kinesin I may coordinate motor activity
J. Biol. Chem.
2004
, vol. 
279
 (pg. 
19201
-
19208
)
103
Schlager
M.
Kapitein
L.C.
Grigor'ev
I.S.
Burzynski
G.
Wulf
P.
Kiejzer
N.
de Graaff
E.
Fukuda
M.
Shepherd
I.
Akhmanova
A.
Hoogenraad
C.C.
Pericentrosomal targeting of Rab6 secretory vesicles by Bicaudal-D-related protein 1 (BICDR-1) regulates neuritogenesis
EMBO J.
2010
, vol. 
29
 (pg. 
1637
-
1651
)
104
Larsen
K.
Xu
J.
Cermelli
S.
Shu
Z.
Gross
S.P.
BicaudalD actively regulates microtubule motor activity in lipid droplet transport
PLoS ONE
2008
, vol. 
3
 pg. 
e3763
 
105
Arimoto
M.
Koushika
S.
Choudhary
B.
Matsumoto
K.
Hisamoto
N.
The Caenorhabditis elegans JIP3 protein UNC-16 functions as an adaptor to link kinesin-1 with cytoplasmic dynein
J. Neurosci.
2011
, vol. 
31
 (pg. 
2216
-
2224
)
106
Horiuchi
D.
Barkus
R.
Pilling
A.
Gassman
A.
Saxton
W.
APLIP1, a kinesin binding JIP-1/JNK scaffold protein, influences the axonal transport of both vesicles and mitochondria in Drosophila
Curr. Biol.
2005
, vol. 
15
 (pg. 
2137
-
2141
)
107
Cavalli
V.
Kujala
P.
Klumperman
J.
Goldstein
L.S.
Sunday Driver links axonal transport to damage signaling
J. Cell Biol.
2005
, vol. 
168
 (pg. 
775
-
787
)
108
Montagnac
G.
Sibarita
J.-B.
Loubéry
S.
Daviet
L.
Romao
M.
Raposo
G.
Chavrier
P.
ARF6 interacts with JIP4 to control a motor switch mechanism regulating endosome traffic in cytokinesis
Curr. Biol.
2009
, vol. 
19
 (pg. 
184
-
195
)
109
Caviston
J.
Holzbaur
E.L.F.
Huntingtin as an essential integrator of intracellular vesicular trafficking
Trends Cell Biol.
2009
, vol. 
19
 (pg. 
147
-
155
)
110
Russo
G.
Louie
K.
Wellington
A.
Macleod
G.
Hu
F.
Panchumarthi
S.
Zinsmaier
K.
Drosophila Miro is required for both anterograde and retrograde axonal mitochondrial transport
J. Neurosci.
2009
, vol. 
29
 (pg. 
5443
-
5455
)
111
Sheets
L.
Ransom
D.
Mellgren
E.
Johnson
S.
Schnapp
B.J.
Zebrafish melanophilin facilitates melanosome dispersion by regulating dynein
Curr. Biol.
2007
, vol. 
17
 (pg. 
1721
-
1734
)
112
Cai
Q.
Lu
L.
Tian
J.-H.
Qiao
H.
Sheng
Z.-H.
Snapin-regulated late endosomal transport is critical for efficient autophagy-lysosomal function in neurons
Neuron
2010
, vol. 
68
 (pg. 
73
-
86
)
113
Bremner
K.
Scherer
J.
Yi
J.
Vershinin
M.
Gross
S.P.
Vallee
R.
Adenovirus transport via direct interaction of cytoplasmic dynein with the viral capsid hexon subunit
Cell Host Microbe
2009
, vol. 
6
 (pg. 
523
-
535
)
114
Bielli
A.
Thörnqvist
P.-O.
Hendrick
A.
Finn
R.
Fitzgerald
K.
McCaffrey
M.
The small GTPase Rab4A interacts with the central region of cytoplasmic dynein light intermediate chain-1
Biochem. Biophys. Res. Commun.
2001
, vol. 
281
 (pg. 
1141
-
1153
)
115
Manneville
J.B.
Jehanno
M.
Etienne-Manneville
S.
Dlg1 binds GKAP to control dynein association with microtubules, centrosome positioning, and cell polarity
J. Cell Biol.
2010
, vol. 
191
 (pg. 
585
-
598
)
116
Navarro
C.
Puthalakath
H.
Adams
J.
Strasser
A.
Lehmann
R.
Egalitarian binds dynein light chain to establish oocyte polarity and maintain oocyte fate
Nat. Cell Biol.
2004
, vol. 
6
 (pg. 
427
-
435
)
117
Fejtova
A.
Davydova
D.
Bichof
F.
Lazarevic
V.
Altrock
W.
Romorini
S.
Schöne
C.
Zuschratter
W.
Kreutz
M.
Garner
C.
, et al. 
Dynein light chain regulates axonal trafficking and synaptic levels of Bassoon
J. Cell Biol.
2009
, vol. 
185
 (pg. 
341
-
355
)
118
Traer
C.
Rutherford
A.
Palmer
K.J.
Wassmer
T.
Oakley
J.
Attar
N.
Carlton
J.
Kremerskothen
J.
Stephens
D.
Cullen
P.
SNX4 coordinates endosomal sorting of TfnR with dynein-mediated transport into the endocytic recycling compartment
Nat. Cell Biol.
2007
, vol. 
9
 (pg. 
1370
-
1380
)
119
Alonso
C.
Miskin
J.
Hernáez
B.
Fernandez-Zapatero
P.
Soto
L.
Canto
C.
Rodriguez-Crespo
I.
Dixon
I.
Escribano
J.
African swine fever virus protein p54 interacts with the microtubular motor complex through direct binding to light-chain dynein
J. Virol.
2001
, vol. 
75
 (pg. 
9819
-
9827
)
120
Wanschers
B.
van de Vorstenbosch
R.A.
Wijers
M.
Wieringa
B.
King
S.M.
Fransen
J.A.
Rab6 family proteins interact with the dynein light chain protein DYNLRB1
Cell Motil. Cytoskeleton
2008
, vol. 
65
 (pg. 
183
-
196
)
121
Watson
P.
Forster
R.
Palmer
K.
Pepperkok
R.
Stephens
D.J.
Coupling of ER exit to microtubules through direct interaction of COPII with dynactin
Nat. Cell Biol.
2005
, vol. 
7
 (pg. 
48
-
55
)
122
Liu
J.-J.
Ding
J.
Kowal
A.
Nardine
T.
Allen
E.
Delcroix
J.-D.
Wu
C.
Mobley
W.
Fuchs
E.
Yang
Y.
BPAG1n4 is essential for retrograde axonal transport in sensory neurons
J. Cell Biol.
2003
, vol. 
163
 (pg. 
223
-
229
)
123
Liu
J.-J.
Ding
J.
Wu
C.
Bhagavatula
P.
Cui
B.
Chu
S.
Mobley
W.
Yang
Y.
Retrolinkin, a membrane protein, plays an important role in retrograde axonal transport
Proc. Natl. Acad. Sci. U.S.A.
2007
, vol. 
104
 (pg. 
2223
-
2228
)
124
Jordens
I.
Fernandez-Borja
M.
Marsman
M.
Dusseljee
S.
Janssen
L.
Calafat
J.
Janssen
H.
Wubbolts
R.
Neefjes
J.
The Rab7 effector protein RILP controls lysosomal transport by inducing the recruitment of dynein–dynactin motors
Curr. Biol.
2001
, vol. 
11
 (pg. 
1680
-
1685
)
125
Wassmer
T.
Attar
N.
Harterink
M.
van Weering
J.R.
Traer
C.J.
Oakley
J.
Goud
B.
Stephens
D.J.
Verkade
P.
Korswagen
H.C.
Cullen
P.J.
The retromer coat complex coordinates endosomal sorting and dynein-mediated transport, with carrier recognition by the trans-Golgi network
Dev. Cell
2009
, vol. 
17
 (pg. 
110
-
122
)
126
Hong
Z.
Yang
Y.
Zhang
C.
Niu
Y.
Li
K.
Zhao
X.
Liu
J.-J.
The retromer component SNX6 interacts with dynactin p150Glued and mediates endosome-to-TGN transport
Cell Res.
2009
, vol. 
19
 (pg. 
1334
-
1349
)