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Dynein promotes sustained axonal growth and Schwann cell remodeling early during peripheral nerve regeneration


Following injury, axons of the peripheral nervous system have retained the capacity for regeneration. While it is well established that injury signals require molecular motors for their transport from the injury site to the nucleus, whether kinesin and dynein motors play additional roles in peripheral nerve regeneration is not well understood. Here we use genetic mutants of motor proteins in a zebrafish peripheral nerve regeneration model to visualize and define in vivo roles for kinesin and dynein. We find that both kinesin-1 and dynein are required for zebrafish peripheral nerve regeneration. While loss of kinesin-1 reduced the overall robustness of axonal regrowth, loss of dynein dramatically impaired axonal regeneration and also reduced injury-induced Schwann cell remodeling. Chimeras between wild type and dynein mutant embryos demonstrate that dynein function in neurons is sufficient to promote axonal regrowth. Finally, by simultaneously monitoring actin and microtubule dynamics in regenerating axons we find that dynein appears dispensable to initiate axonal regrowth, but is critical to stabilize microtubules, thereby sustaining axonal regeneration. These results reveal two previously unappreciated roles for dynein during peripheral nerve regeneration, initiating injury induced Schwann cell remodeling and stabilizing axonal microtubules to sustain axonal regrowth.

Author summary

Nerve regeneration requires coordinated responses from multiple cell types after injury. Axons must extend from the neuronal cell body back towards their targets, while surrounding Schwann cells enter a repair cell state in which they promote regeneration. While nerves of the peripheral nervous system can regrow, it is estimated that fewer than 10 percent of patients fully recover function after nerve injury. In order to understand the mechanisms by which peripheral nerves regrow, we used live cell imaging in the zebrafish to observe the process of nerve regeneration, monitoring axons and Schwann cells simultaneously during this process. Using genetic mutants, we identified a role for the molecular motors kinesin-1 and dynein in promoting axonal regrowth. Furthermore, we found that dynein plays an additional role in Schwann cell response to injury. Thus, we demonstrate that molecular motors are required in multiple cell types to promote nerve regeneration.


Axons of the mature peripheral nervous system have retained a remarkable ability for regeneration. Although simple in concept, peripheral nerve regeneration is a complex process that requires extrinsic as well as intrinsic mechanisms. Chief amongst the intracellular mechanisms that contribute to axonal regeneration are microtubule organization and dynamics as well as axonal transport. It has long been known that following injury the pool of dynamic microtubules at the lesion site, as well as axonal transport, increase [13]. Given the central role of both microtubule dynamics and axonal transport in promoting axonal regeneration, factors that regulate both processes are prime candidates for regulating peripheral nerve regeneration.

The molecular motor proteins kinesin-1 and dynein are key regulators of both microtubule organization and axonal transport and have both been implicated in peripheral nerve regeneration. Kinesin-1 is an anterograde motor that is essential for maintaining neuronal homeostasis by transporting cargos, including organelles and mRNA, from the cell body toward synaptic terminals. Kinesin-1 has also been shown to drive axonal outgrowth during development and after injury [4,5]. Dynein has similarly been studied for its role in maintaining homeostasis by transporting cargo, however dynein moves cargo retrogradely towards the cell body. Dynein also plays an important role in axonal injury by trafficking injury signals, including components of JNK and ERK MAPK pathways, which are generated at the lesion site and actively transported to the cell body [6,7]. There these injury signals initiate a regenerative response, characterized first by upregulation of regeneration-associated genes that prevent neuronal cell death, and by initiating a genetic program that promotes regrowth of injured axons back to their original targets [8,9].

More recently it has become clear that in addition to its role in retrograde transport, dynein also functions in cytoskeletal organization and maintenance. For example, in C. elegans dynein regulates local microtubule dynamics in dendrites to promote microtubule stabilization [10]. Additionally, in the axon dynein transports microtubules to establish and maintain microtubule polarity [1113]. Finally, besides its preeminent role in axonal homeostasis, dynein is also required for Schwann cell development and myelination [14]. Yet despite dynein’s well documented roles in both axons and glial cells, the effects of dynein on the cellular behaviors of regenerating axons and their associated glial cells in intact animals have not been examined.

In order to examine the diverse cellular functions of molecular motors in multiple cell types, we combined genetic mutants with live imaging of nerve regeneration in larval zebrafish, as previously described [15]. This allowed us to study the real-time dynamics of regenerating axons and surrounding Schwann cells in a whole organism context. We find that the molecular motors kinesin-1 and dynein, albeit to different degrees, are both required for axonal regrowth in vivo. Furthermore, we find that dynein is also required to initiate injury-induced morphology changes in Schwann cells, however wild type neurons transplanted into otherwise dynein mutant animals are able to regrow robustly, indicating that neuronal dynein is sufficient to promote axonal regrowth. Finally, we find that dynein is dispensable for initiation of axonal regrowth but is required to stabilize microtubules in injured axons to generate persistent, long-range regrowth. These findings elucidate previously unknown roles for dynein in the initiation of injury-induced Schwann cell behaviors, and identify a distinct role for dynein in promoting axonal regeneration through persistent axonal regrowth via microtubule stabilization.


Kinesin-1 and dynein are critical for peripheral nerve regeneration in vivo

To determine the in vivo roles of molecular motors in peripheral nerve regeneration, we first assessed regeneration in mutants lacking kif5aa, which encodes the neuron-specific Kif5A heavy chain of the conventional anterograde motor Kinesin-1. We have previously shown that laser mediated transection of motor nerves in larval zebrafish initiates a Schwann cell dependent peripheral nerve regeneration program reminiscent of what is observed in adult vertebrates [16]. Following their complete transection at 5 days post-fertilization (dpf), ventral motor nerves exhibit Schwann cell dependent functional regeneration by 48 hours post-transection (hpt) [15] (Fig 1A and 1B). Prior to transection, kif5aa-/- motor nerves were indistinguishable from wild type nerves (Fig 1C). By 48 hpt, motor axons in kif5aa-/- mutants had regrown across the full extent of the ventral myotome, although when compared to wild type siblings the number of fascicles that reached their ventral targets was reduced (Fig 1D and 1E). Using a previously established semi-quantitative scoring index (for details see Materials and methods and [15] we confirmed that compared to wild type siblings, motor axons in kif5aa mutants exhibited reduced regeneration (p = 0.0487, Fisher’s exact test).

Fig 1. Dynein and Kinesin-1 are required for peripheral nerve regeneration in vivo.

(A) Wild type motor nerve pre-lesion (red box, transection site; scale bar = 20 μm). (B) By 48 hpt, several fascicles have regrown fully across the ventral myotome (green arrowheads, regrown axons, strong regeneration). (C) kif5aa-/- motor nerve pre-lesion. (D) At 48 hpt, some axons have extended across the myotome (blue arrowheads, regrown axons, moderate regeneration). (E) Quantification of kif5aa mutant regeneration at 48 hpt (wild type siblings, n = 66 nerves; kif5aa-/-, n = 20 nerves, p = 0.0487, Fisher’s exact test). (F) dync1h1-/- motor nerve pre-lesion. (G) By 48 hpt, regrowing axons have extended slightly but failed to reach the ventral extend of the myotome (red arrowheads, stalled axons, no/weak regeneration). (H) Quantification of dync1h1 mutant regeneration at 48 hpt (dync1h1+/+, n = 59 nerves; dync1h1+/-, n = 21 nerves; dync1h1-/-, n = 25 nerves; p = 0.007; p = 0.0006; p<0.0001, respectively, Fisher’s exact test).

We next assessed motor axon regeneration in genetic mutants for the dynein heavy chain gene (dync1h1) which encodes a core component of the retrograde motor dynein. Prior to injury at 5 dpf, dync1h1-/- motor axons exhibit normal architecture, presumably due to the large maternal load sufficient to promote axonal development [17] (Fig 1F). In contrast, following transection, motor axons in dync1h1-/- mutant animals frequently failed to extend beyond the transection site (Fig 1G, quantified in Fig 1H). Analysis of dynein heterozygotes revealed a less severe, although still significant, defect in axonal regrowth, demonstrating a dose-dependent requirement for dynein in promoting axonal regrowth. The severity of the regeneration phenotype we observed in homozygous dync1h1-/- mutants was significantly stronger than that present in kif5aa-/- mutants. This is consistent with the notion that other heavy chains of Kinesin-1 as well as other Kinesin family motors might compensate for the absence of kif5aa [18,19]. In contrast, dynein is the sole protein responsible for microtubule-associated retrograde transport, and therefore the regeneration phenotype we observe in dync1h1-/- mutants likely represents a complete block of retrograde transport. We therefore focused on further defining the role of dynein in peripheral nerve regeneration.

Dynein is required for injury-induced Schwann cell remodeling

In addition to its important and well-studied function in neurons, dynein is also required for proper differentiation and myelination of Schwann cells during development [14]. Furthermore, in zebrafish lacking Schwann cells, regenerating axons sprout from the proximal nerve stump but fail to grow across the injury gap [20], somewhat reminiscent of the phenotype we observe in dynein mutants. Given the importance of Schwann cells for peripheral nerve regeneration and the role of dynein in Schwann cell development, we sought to determine whether dynein is also required for the Schwann cell response to injury, characterized by stereotyped changes in Schwann cell morphology.

We have previously shown that before injury, Schwann cell membranes ensheathe individual motor axons, and that following nerve transection when axons fragment, Schwann cell membranes reorganize, changing from a smooth, tube-like appearance to a more rounded and granular morphology [20], indicative of their transition to an activated, dedifferentiated state, known as the repair cell state that promotes axonal regeneration. Previous studies revealed that in dynein mutants, Schwann cells developmentally arrest at the promyelinating stage [14]. We therefore first wanted to determine whether immature Schwann cells are able to respond appropriately to injury. For this we examined a mutant for the G-protein coupled receptor GPR126, in which Schwann cells also arrest at the promyelinating stage [21], similar to what has been reported for dync1h1 mutants. Importantly, in contrast to dynein mutants, gpr126 mutants do not exhibit an obvious deficit in axonal regeneration (n = 37 nerves from 12 +/+ or +/- wild type larvae and 33 nerves from 11 gpr126-/- mutants, respectively; p>0.85, Fisher’s exact test). Analysis of Schwann cells dynamics in gpr126 mutants revealed that Schwann cells respond to injury by extending their membranes dramatically compared to their pre-injury state, indistinguishable from wild type Schwann cells (Fig 2A–2D). This demonstrates that developmentally arrested Schwann cells are still able to respond appropriately to nerve injury.

Fig 2. Dynein is required for injury-induced Schwann cell morphology changes.

(A-F) Schwann cells in 5 dpf larvae labeled by Tg(sox10:mRFP). (A) Prior to injury, wild type Schwann cells have smooth, straight membranes that are tightly associated with the axonal track (scale bar = 5 μm). (B) After axonal fragmentation, Schwann cell membranes change morphology and widen to accommodate axonal debris. (C) Prior to injury, gpr126-/- Schwann cells are loosely associated with axons as they do not myelinate. (D) After axonal fragmentation, gpr126-/- Schwann cells are able to change morphology and widen. (E) Prior to injury, dync1h1-/- Schwann cells are loosely associated with axons as they also do not myelinate. (F) After axonal fragmentation, dync1h1-/- Schwann cell membranes maintain an elongated conformation and do not dramatically change morphology, indicating a disrupted injury response. (G) Quantification of Schwann cell width pre- and post-fragmentation in gpr126 and dync1h1 mutants.

Having determined that promyelinating Schwann cells are competent to respond appropriately to nerve injury, we next examined the behavior of dync1h1-/- mutant Schwann cells. Unlike wild type and gpr126 mutant Schwann cells, we find that following nerve transection dync1h1-/- mutant Schwann cells fail to initiate any morphological changes, and instead retain their pre-injury morphology and membrane position for the duration of the imaging period (up to five hours), arguing against a delay in onset but rather for a complete lack of Schwann cell injury response (Fig 2E and 2F). To quantify this phenotype, we measured the changes in Schwann cell width following nerve transection as a simpler proxy for the complex changes in Schwann cell morphology (Fig 2G). This revealed that while wild type and gpr126-/- Schwann cells significantly increase in width after injury, dync1h1-/- Schwann cells show no significant change. Thus, while dync1h1-/- mutant axons initiate fragmentation following injury, their associated Schwann cells fail to respond, consistent with the idea that dynein is critical for injury-induced Schwann cell remodeling.

Neuronal dynein is sufficient to promote axonal regrowth

Our results reveal injury-induced phenotypes in two cell types after injury in dynein mutants, and we therefore wondered whether dynein functions in neurons or Schwann cells to promote axonal regrowth. To determine the cell type in which dynein functions to promote axonal regrowth, we generated chimeras at the blastula stage [22] that contained wild type motor neurons and axons in otherwise dync1h1-/- larvae (Fig 3A and 3B). Control transplantations have previously shown that wild type cells transplanted into wild type embryos generate motor neurons that are morphologically and functionally unaffected by transplantation [23]. Following development and subsequent transection in a dync1h1-/- environment, wild type axons were able to regenerate robustly for the first 9 hours after sprouting (Fig 3C–3F), in a manner indistinguishable from wild type axons in a fully wild type environment. This indicates that restoring dynein specifically in neurons in a dynein mutant is sufficient to promote axonal regrowth, demonstrating a neuron-intrinsic role for dynein during peripheral nerve regeneration.

Fig 3. Neuronal dynein is sufficient to promote axonal regrowth.

(A) ~10 rhodamine-labeled cells were transplanted from wild type blastulas to dync1h1-/- blastulas. (B) At 5 dpf, nerves contained wild type neurons (transplanted cells labeled by rhodamine-dextran, magenta) in a dync1h1-/- larva (host motor neurons labeled by Tg(mnx1:GFP), green; scale bar = 10 μm). (C-E) After transection, wild type axons (magenta arrowheads) are able to regrow robustly in the dync1h1-/- embryo, while dync1h1-/- host axons regrow significantly less (green arrowheads; scale bar = 10 μm). (F) Quantification of growth cone displacement in dync1h1-/- host axons and transplanted wild type axons. Open circles indicate dync1h1-/- mutant axons that grew along transplanted wild type axons. (G-I) Some dync1h1-/- axons demonstrated improved regeneration in the presence of wild type axons in the same nerve. Here, a dync1h1-/- axon (green arrowheads) follows along a previously regrown wild type axon (magenta arrowheads; scale bar = 5 μm).

Interestingly, we found that dync1h1-/- axons that had wild type axons in the same nerve regrew more robustly than dync1h1-/- axons in nerves with no transplanted cells (14.23 ± 2.06 μm growth in dync1h1-/- larvae without transplants, see below; 39.33 ± 4.72 μm growth in dync1h1-/- larvae with transplants, Fig 3F). In several instances, we observed dync1h1-/- axons growing along previously extended wild type axons (Fig 3G–3I). This indicates that the presence of wild type axonal regrowth is able to partially rescue the dync1h1-/- axonal regrowth defects, likely through cell-cell adhesions between the dynein mutant axon and wild type axons.

Dynein stabilizes axonal growth during regeneration

We next asked how dynein promotes axonal regeneration within peripheral nerves. Peripheral nerve regeneration is a dynamic process composed of several defined stages, starting with growth cones emerging from the proximal stump and starting to probe the injury gap environment. This is followed by stabilization of axonal regrowth across the injury gap and along the correct trajectory, and finally rapid, sustained axonal regrowth towards their original targets [24]. We used live cell imaging after nerve transection to quantify axonal dynamics in dynein mutants and determine which of these stages require dynein. In wild type siblings, we observed growth cones emerging from the proximal stump extending (3.54 events per 8 hours) and retracting (1.08 events per 8 hours) repeatedly, consistent with the idea that these growth cones are probing the injury gap for a path towards their original targets (Fig 4A and 4B). We found that dync1h1-/- axons exhibit similar frequencies of axonal extensions and retractions (Fig 4C and 4D), suggesting that they probe the injury gap as actively as their wild type siblings (Fig 4E).

Fig 4. Dynein stabilizes axonal extensions during regeneration.

(A-B) In wild type animals, regenerating axons begin probing the environment by extending and retracting (green and red arrowheads, respectively; scale bar = 5 μm). (C-D) dync1h1-/- axons also extend and retract after injury. (E) Quantification of extension and retraction events in wild type siblings (n = 13 axons) and dync1h1-/- axons (n = 13 axons). (F-G) Measurement of overall growth cone displacement from transection site ~16 hpt in wild type siblings (F; blue arrowheads, growth cones; scale bar = 10 μm) and dync1h1-/- (G; red arrowheads, growth cones). (H) Quantification of growth cone displacement ~16 hpt (wild type siblings, n = 15; dync1h1-/-, n = 10; p = 0.0005, unpaired t-test).

We next examined the second stage of axonal regeneration when axons become stabilized and then extend toward their original targets. To quantify this process we measured the overall displacement of regenerating growth cones over the first ~8 hours after sprouting began. We found that the majority of regenerating wild type axons grew beyond the transection site within 8 hours of sprouting (Fig 4F), travelling an average of 41.49 μm (SEM ± 5.84) over this time period. In contrast, regenerating dync1h1-/- axons rarely extended beyond the transection site (Fig 4G), travelling an average of 14.23 μm (SEM ± 2.06) and never exceeding 21.94 μm in growth. Moreover, quantification of growth cone displacement at 8 hours post transection revealed that compared to regenerating wild type axons, dync1h1-/- axons exhibited a significant decrease in axonal extension (Fig 4H). Combined these results argue that rather than initiating growth cone sprouting and short range axonal extensions, dynein predominantly acts early during axonal regeneration to stabilize regenerating axons thereby promoting persistent, long-range regrowth.

Dynein stabilizes microtubules to promote persistent regrowth

Dynein has recently been shown to play a critical role in generating and maintaining microtubule organization, both processes central to axonal growth [10,11,25,26]. To determine whether dynein regulates microtubule dynamics in axons during regeneration, we used a transgenic line that simultaneously labels actin and microtubules in motor neurons (mnx1:Gal4; UAS:lifeact-GFP-v2a-EB3-RFP). Growth cone extension occurs in three stages: first, protrusion driven by F-actin, then engorgement driven by microtubule-based transport of organelles and vesicles, and finally consolidation in which the growth cone contracts and stabilizes to form a cylindrical axon shaft [27]. In regenerating wild type axons, filopodia extend at the growth cone and microtubules follow behind, stabilizing and consolidating newly formed protrusions (Fig 5A–5D). The majority of regenerating dync1h1-/- axons (n = 30/37) displayed one of two phenotypes characteristic for microtubule disruption. In 59 percent (n = 22/37) we observed filopodia extension followed briefly by microtubule extension (Fig 5E and 5F) and then arrest at the engorgement stage before finally retracting (Fig 5G and 5H). In 22 percent (n = 8/37) of regenerating dync1h1-/- axons, microtubules faithfully followed filopodia extending at growth cones. However, rather than consolidating in the proximal growth cone, they adopted a looped conformation at the leading edge of the growth cone, leading to stalling and retraction (Fig 5I and 5M). The remaining 19 percent had straight, ordered microtubules (n = 7/37). This suggests that a lack of dynein may lead to loss of microtubule organization at regenerating growth cones and stalling of regenerating axons early during the regeneration process. To determine whether stabilizing microtubules during axonal regrowth could compensate for a lack of dynein, we transected nerves in dynein mutant larvae and subsequently treated the larvae with taxol. We used timelapse imaging to assess axonal regrowth dynamics and found that in dynein mutant embryos stabilizing microtubules with 5 μm taxol partially rescued axonal regrowth (Fig 5N). Combined, these findings support a model by which dynein plays a critical role in regulating microtubule dynamics, thereby stabilizing growth of regenerating axons as they initiate their trajectory across the injury gap and towards their original targets. Thus, we demonstrate a role for dynein in promoting axonal extension via microtubule stabilization, as well as a previously uncharacterized role in initiating Schwann cell response to injury.

Fig 5. Dynein stabilizes microtubules to promote persistent regrowth.

(A-D) Regenerating wild type axons first extend actin protrusions then extended microtubules, leading to stable growth (scale bar = 5 μm; green arrowheads, actin; magenta arrowheads, microtubules). (E-H) dync1h1-/- axons extend actin protrusions followed by microtubule growth that arrests during growth cone engorgement and leads to axon retraction (G,H). (I-L) dync1h1-/- axons extend actin protrusions but microtubules form aberrant loop structures (magenta arrowheads), preventing further regrowth. (M) Quantification of microtubule organization in regrowing axons of dync1h1 mutants (siblings, n = 19 axons; dync1h1-/-, n = 37 axons; p<0.0001, Fisher’s exact test). (N) Quantification of growth cone displacement ~12 hpt with and without taxol treatment (dync1h1-/- with DMSO, n = 8, dync1h1-/- with 5 μM taxol, n = 12, p = 0.0159; wild type siblings with DMSO, n = 4, wild type siblings with 5 μM taxol, n = 4, p = 0.0571).

Materials and methods

Ethics statement

All experiments were conducted according to an Animal Protocol fully approved by the Uni- versity of Pennsylvania Institutional Animal Care and Use Committee (IACUC) on January 24, 2014, protocol number 803446. Veterinary care is under the supervision of the University Laboratory Animal Resources (ULAR) of the University of Pennsylvania.

Zebrafish genetics and transgenes

All transgenic lines were maintained in the Tübigen or Tupfel long fin genetic background and raised as previously described [22]. The Tg(mnx1:GFP)ml2 line [28] was used to label spinal motor nerves and the Tg(sox10(7.2):mRFP)vu234 line [29] was used to label Schwann cells. The Tg(UAS:lifeact-GFP-v2a-EB3-RFP) line was used to label microtubules and actin. The dync1h1hi3684Tg [30] and gpr126stl47 [31] mutant strains were used and genotyping protocols were performed as previously described.

Nerve transection and live imaging

Nerve transection and live imaging were performed as previously described [15].

Axon growth extent quantification

Axon growth extent quantification was performed as previously described [20]. Transected nerves in which axons failed to regrow or did not extend through the entire length of the ventral myotome are categorized as “no/weak regeneration.” Nerves with at least one fascicle that extended through the entire length of the ventral myo- tome are categorized as “moderate regeneration.” Finally, nerves with two or more fascicles extending through the entire length of the ventral myotome are categorized as “strong regeneration.”

Axon extension and retraction quantification

Axons were imaged every 15 minutes from ~7 to ~16 hpt. Extensions and retractions were defined as growth or retraction of >1 μm between timelapse frames and number of extension and retraction events was counted. Continued movements of the same direction in a subsequent frame were not counted as new events. Measurements were performed on each visibly distinct axon in a nerve.

Growth cone displacement quantification

Axons imaged at ~16 hpt were measured by drawing a line from the spinal cord exit point to the growth cone. Measurements were performed on each visibly distinct axon in a nerve.

Schwann cell width quantification

Axons and Schwann cells were imaged before transection and every 15 minutes from ~1 to ~5 hpt. Schwann cell width was measured at the widest point in pre- and post-transection images. Using ImageJ, a line was drawn from one edge of the Schwann cell membrane to the other in an orientation perpendicular to the motor nerve and was measured in microns.

Cell transplantation for chimera analysis

Cell transplantations were performed as previously described [32]. Wild type cells were transplanted into dync1h1-/- embryos in areas known to develop into motor neurons. Larvae were screened at 5 dpf to identify nerves that contained transplanted motor neurons and no other transplanted cell types along the path of the ventral motor nerve. Transection, imaging, and quantification of growth cone displacement in identified nerves were performed as described above.

Taxol treatment protocol

Larvae were transected according to the above protocol. 3 hours after transection, embryos were bathed in embryo medium with either 1% DMSO or 1% DMSO and 5 μM taxol (paclitaxel, Life Technologies #P3456). When mounted for overnight timelapse imaging, larvae were bathed in Ringer’s solution with either 1% DMSO or 1% DMSO and 5 μM taxol for the duration of imaging.

Image processing

Image stacks were compressed into maximum intensity projections (MIPs) in Slidebook 6 then processed using ImageJ and Photoshop to normalize brightness and contrast.

Statistical analysis

Fisher’s exact and Student’s t tests were performed on all applicable datasets.


Dynein stabilizes axonal extensions during regeneration

Nerve injury induces a local signaling cascade that leads to the production of axon intrinsic signals at the lesion site [33]. There is overwhelming evidence that dynein is critical to transport these injury signals from the lesion site to the cell body where they initiate a neuronal injury response [3436]. We find that in presumptive dynein null mutants, injured neurons robustly respond to the injury and within ~8–10 hours, regenerating axons sprout from the proximal stump, indistinguishable from what we observe in wild type animals. This raises the question whether axonal sprouting can occur independently of dynein-dependent injury induced signals, or whether in our zebrafish model dynein-mediated retrograde transport is less important to mount an injury response? One clear difference between rodent models and our model is the distance between the injury site and the neuronal cell bodies. In rodent sciatic nerve models lesions are introduced millimeters away from neuronal cell bodies [36], while in larval zebrafish—due to the smaller animal size—lesions are generated about 10–50 μm away from neuronal cell bodies [15] Thus, it is conceivable that due to the almost 100-fold reduction in distance between lesion site and cell body, injury signal propagation from the lesion site to the cell body is less dependent on dynein function. Although it remains unclear how injury signal propagation can occur independent of dynein, this provided us with the unique opportunity to examine dynein’s role in peripheral nerve regeneration beyond its role in injury signal transport.

Endpoint analyses at 48 hpt uncovered a clear role for dynein in peripheral nerve regeneration, with clear effects on both axonal regrowth as well as on injury-induced Schwann cell remodeling (Figs 1, 2 and 4). Using live-imaging to visualize the early stages of the regeneration process, we found that dynein promotes the stabilization and growth of long-range axonal projections, providing compelling evidence that apart from its well-documented role in retrograde injury signal transport, dynein also plays a critical role in sustaining axonal regrowth. Moreover, simultaneously visualizing the cellular behavior of both axons and Schwann cells revealed that loss of dynein prevented injury-induced Schwann cell remodeling. The transition of Schwann cells from their fully differentiated state to a repair cell state is a well-documented and integral aspect of peripheral nerve regeneration [37,38], accompanied by dramatic morphological changes to the Schwann cell, as the cell breaks down its myelin and extends its membrane to engulf axonal debris [39,40]. Dynein regulates several steps of membrane trafficking, including ER to golgi transport, as well as endosomal trafficking [41], so it is conceivable that dynein plays a direct, cell-autonomous role in this process. Alternatively, the inability of Schwann cells to initiate the remodeling process might be a consequence of strongly reduced axonal regrowth, and future experiments will be required to test a possible Schwann cell-specific role for dynein in the remodeling process.

Given that dynein mutants exhibit defects in axonal regrowth and Schwann cell morphology, we performed chimeric analysis experiments. These experiments revealed that dynein function in injured neurons is sufficient to sustain axonal regeneration. Importantly in our chimera experiments, of the roughly 60 axons contributing to an individual motor nerve [42], on average only 1–3 transplanted wild type axons were present. This low level of chimerism was critical to evaluate regrowth capacity of individual wild type axons. This also revealed that the presence of individual wild type axonal regrowth facilitated regrowth of individual, neighboring dynein deficient axons (Fig 5G–5I). At the same time, the low level of chimerism precluded us from asking whether neuronal dynein restored all aspects of peripheral nerve regeneration, including the overall robustness of axonal regrowth for a whole nerve and injury-induced Schwann cell remodeling. Thus, while neuronal dynein plays a critical role in sustaining axonal regrowth, we cannot exclude the possibility that dynein function in Schwann cells also contributes to peripheral nerve regeneration.

Dynein promotes axonal regeneration by modulating microtubule dynamics

Cytoskeletal dynamics are critical to growth cone formation [43], axonal outgrowth during development [44], and axonal regeneration [45]. Previous studies have revealed that microtubule stabilization promotes axonal regrowth after injury both in vitro and in vivo [4648]. Interestingly, studies of C. elegans dynein heavy chain mutants recently revealed that dynein acts locally in dendrites to stabilize microtubules [10]. This raised the possibility that dynein may also act locally in regenerating axons to stabilize microtubules. We assessed cytoskeletal dynamics during regeneration using a transgene that allowed us to visualize actin and microtubules simultaneously in live, regenerating axons. This revealed that while actin dynamics were grossly unaffected in dynein mutant axons, microtubules often appeared unstable and disordered, with some axons exhibiting looping microtubule configurations reminiscent of those seen in the dendrites of C. elegans dynein heavy chain mutants [10]. Thus, our results provide compelling evidence that besides its well-documented role in retrograde transport, dynein also promotes microtubule stability critical for growth cone advancement [49], providing a potential mechanism for the rapid and sustained extension observed during wild type axonal regrowth, and deficient in dynein mutants (Fig 2).

Dynein is also known to modulate microtubule dynamics is through microtubule sliding [50], providing an alternative mechanism. This might be a direct effect or may affect microtubule sliding indirectly via modulation of kinesin-1 as these motors have been shown to transport each other directly with one another [51]. Taken together, our results suggest that beyond its function in retrograde injury signaling dynein has a multifaceted role in nerve regeneration that warrants further studies.

Supporting information

S1 Movie. Wild type axon regrowth in a dynein mutant background.


S3 Movie. Dynein mutant axon regrowth dynamics.


S4 Movie. Wild type cytoskeletal dynamics during axonal regrowth.


S5 Movie. Dynein mutant cytoskeletal dynamics during axonal regrowth (stalling).


S6 Movie. Dynein mutant cytoskeletal dynamics during axonal regrowth (looping).



We would like to thank Dr. Monk (OHSU), Dr. Marlow (MSSM) for reagents and members of the Granato lab for discussion and comments.


  1. 1. Oblinger MM, Lasek RJ. Axotomy-induced alterations in the synthesis and transport of neurofilaments and microtubules in dorsal root ganglion cells. Journal of Neuroscience. 1988;8: 1747–1758. pmid:3130470
  2. 2. McQuarrie IG, Jacob JM. Conditioning nerve crush accelerates cytoskeletal protein transport in sprouts that form after a subsequent crush. J Comp Neurol. Wiley-Blackwell; 1991;305: 139–147. pmid:1709646
  3. 3. Misgeld T, Kerschensteiner M, Bareyre FM, Burgess RW, Lichtman JW. Imaging axonal transport of mitochondria in vivo. Nat Meth. 2007;4: 559–561. pmid:17558414
  4. 4. Winding M, Kelliher MT, Lu W, Wildonger J, Gelfand VI. Role of kinesin-1-based microtubule sliding in Drosophila nervous system development. Proceedings of the National Academy of Sciences. National Academy of Sciences; 2016;113: E4985–94. pmid:27512046
  5. 5. Lu W, Lakonishok M, Gelfand VI. Kinesin-1-powered microtubule sliding initiates axonal regeneration in Drosophila cultured neurons. Fehon R, editor. Mol Biol Cell. 2015;26: 1296–1307. pmid:25657321
  6. 6. Cavalli V. Sunday Driver links axonal transport to damage signaling. The Journal of Cell Biology. 2005;168: 775–787. pmid:15738268
  7. 7. Perlson E, Hanz S, Ben-Yaakov K, Segal-Ruder Y, Seger R, Fainzilber M. Vimentin-Dependent Spatial Translocation of an Activated MAP Kinase in Injured Nerve. Neuron. 2005;45: 715–726. pmid:15748847
  8. 8. Abe N, Cavalli V. Nerve injury signaling. Curr Opin Neurobiol. 2008;18: 276–283. pmid:18655834
  9. 9. Hanz S, Fainzilber M. Retrograde signaling in injured nerve? the axon reaction revisited. J Neurochem. 2006;99: 13–19. pmid:16899067
  10. 10. Yogev S, Maeder CI, Cooper R, Horowitz M, Hendricks AG, Shen K. Local inhibition of microtubule dynamics by dynein is required for neuronal cargo distribution. Nat Commun. Nature Publishing Group; 2017;8: 1–14. pmid:28406181
  11. 11. Ahmad FJ, Echeverri CJ, Vallee RB, Baas PW. Cytoplasmic dynein and dynactin are required for the transport of microtubules into the axon. The Journal of Cell Biology. 1998;140: 391–401. pmid:9442114
  12. 12. Zheng Y, Wildonger J, Ye B, Zhang Y, Kita A, Younger SH, et al. Dynein is required for polarized dendritic transport and uniform microtubule orientation in axons. Nat Cell Biol. 2008;10: 1172–1180. pmid:18758451
  13. 13. Rao AN, Patil A, Black MM, Craig EM, Myers KA, Yeung HT, et al. Cytoplasmic Dynein Transports Axonal Microtubules in a Polarity-Sorting Manner. CellReports. ElsevierCompany; 2017;19: 2210–2219. pmid:28614709
  14. 14. Langworthy MM, Appel B. Schwann cell myelination requires Dynein function. Neural Dev. 2012;7: 37. pmid:23167977
  15. 15. Rosenberg AF, Wolman MA, Franzini-Armstrong C, Granato M. In vivo nerve-macrophage interactions following peripheral nerve injury. Journal of Neuroscience. 2012;32: 3898–3909. pmid:22423110
  16. 16. Rosenberg AF, Isaacman-Beck J, Franzini-Armstrong C, Granato M. Schwann cells and deleted in colorectal carcinoma direct regenerating motor axons towards their original path. J Neurosci. Society for Neuroscience; 2014;34: 14668–14681. pmid:25355219
  17. 17. Insinna C, Baye LM, Amsterdam A, Besharse JC, Link BA. Analysis of a zebrafish dync1h1 mutant reveals multiple functions for cytoplasmic dynein 1 during retinal photoreceptor development. Neural Dev. 2010;5: 12. pmid:20412557
  18. 18. Miki H, Setou M, Kaneshiro K, Hirokawa N. All kinesin superfamily protein, KIF, genes in mouse and human. Proceedings of the National Academy of Sciences. National Academy of Sciences; 2001;98: 7004–7011. pmid:11416179
  19. 19. Campbell PD, Shen K, Sapio MR, Glenn TD, Talbot WS, Marlow FL. Unique function of Kinesin Kif5A in localization of mitochondria in axons. J Neurosci. Society for Neuroscience; 2014;34: 14717–14732. pmid:25355224
  20. 20. Rosenberg AF, Isaacman-Beck J, Franzini-Armstrong C, Granato M. Schwann cells and deleted in colorectal carcinoma direct regenerating motor axons towards their original path. J Neurosci. Society for Neuroscience; 2014;34: 14668–14681. pmid:25355219
  21. 21. Monk KR, Naylor SG, Glenn TD, Mercurio S, Perlin JR, Dominguez C, et al. A G protein-coupled receptor is essential for Schwann cells to initiate myelination. Science. 2009;325: 1402–1405. pmid:19745155
  22. 22. Mullins MC, Hammerschmidt M, Haffter P, Nusslein-Volhard C. Large-scale mutagenesis in the zebrafish: in search of genes controlling development in a vertebrate. Curr Biol. 1994;4: 189–202. pmid:7922324
  23. 23. Eisen JS. Determination of primary motoneuron identity in developing zebrafish embryos. Science. 1991;252: 569–572. pmid:1708527
  24. 24. Isaacman-Beck J, Schneider V, Franzini-Armstrong C, Granato M. The lh3 Glycosyltransferase Directs Target-Selective Peripheral Nerve Regeneration. Neuron. 2015;88: 691–703. pmid:26549330
  25. 25. del Castillo U, Winding M, Lu W, Gelfand VI. Interplay between kinesin-1 and cortical dynein during axonal outgrowth and microtubule organization in Drosophila neurons. Elife. eLife Sciences Publications Limited; 2015;4: e10140. pmid:26615019
  26. 26. Hendricks AG, Lazarus JE, Perlson E, Gardner MK, Odde DJ, Goldman YE, et al. Dynein tethers and stabilizes dynamic microtubule plus ends. Curr Biol. 2012;22: 632–637. pmid:22445300
  27. 27. Goldberg DJ, Burmeister DW. Stages in axon formation: observations of growth of Aplysia axons in culture using video-enhanced contrast-differential interference contrast microscopy. The Journal of Cell Biology. 1986;103: 1921–1931. pmid:3782290
  28. 28. Flanagan-Steet H, Fox MA, Meyer D, Sanes JR. Neuromuscular synapses can form in vivo by incorporation of initially aneural postsynaptic specializations. Development. Oxford University Press for The Company of Biologists Limited; 2005;132: 4471–4481. pmid:16162647
  29. 29. Kucenas S, Takada N, Park H-C, Woodruff E, Broadie K, Appel B. CNS-derived glia ensheath peripheral nerves and mediate motor root development. Nat Neurosci. 2008;11: 143–151. pmid:18176560
  30. 30. Amsterdam A, Nissen RM, Sun Z, Swindell EC, Farrington S, Hopkins N. Identification of 315 genes essential for early zebrafish development. Proc Natl Acad Sci USA. 2004;101: 12792–12797. pmid:15256591
  31. 31. Petersen SC, Luo R, Liebscher I, Giera S, Jeong S-J, Mogha A, et al. The Adhesion GPCR GPR126 Has Distinct, Domain-Dependent Functions in Schwann Cell Development Mediated by Interaction with Laminin-211. Neuron. 2015;85: 755–769. pmid:25695270
  32. 32. Carmany-Rampey A, Moens CB. Modern mosaic analysis in the zebrafish. Methods. 2006;39: 228–238. pmid:16829130
  33. 33. Zhang XP, Ambron RT. Positive injury signals induce growth and prolong survival in Aplysia neurons. J Neurobiol. 2000;45: 84–94. pmid:11018770
  34. 34. Schmied R, Ambron RT. A nuclear localization signal targets proteins to the retrograde transport system, thereby evading uptake into organelles in aplysia axons. J Neurobiol. 1997;33: 151–160. pmid:9240371
  35. 35. Wellmann H, Kaltschmidt B, Kaltschmidt C. Retrograde transport of transcription factor NF-kappa B in living neurons. J Biol Chem. American Society for Biochemistry and Molecular Biology; 2001;276: 11821–11829. pmid:11096106
  36. 36. Hanz S, Perlson E, Willis D, Zheng J-Q, Massarwa R, Huerta JJ, et al. Axoplasmic importins enable retrograde injury signaling in lesioned nerve. Neuron. 2003;40: 1095–1104. pmid:14687545
  37. 37. Arthur-Farraj PJ, Latouche M, Wilton DK, Quintes S, Chabrol E, Banerjee A, et al. c-Jun Reprograms Schwann Cells of Injured Nerves to Generate a Repair Cell Essential for Regeneration. Neuron. 2012;75: 633–647. pmid:22920255
  38. 38. Fontana X, Hristova M, Da Costa C, Patodia S, Thei L, Makwana M, et al. c-Jun in Schwann cells promotes axonal regeneration and motoneuron survival via paracrine signaling. The Journal of Cell Biology. 2012;198: 127–141. pmid:22753894
  39. 39. Glenn TD, Talbot WS. Signals regulating myelination in peripheral nerves and the Schwann cell response to injury. Curr Opin Neurobiol. 2013. pmid:23896313
  40. 40. Zochodne DW. The challenges and beauty of peripheral nerve regrowth. Journal of the peripheral nervous system: JPNS. Wiley/Blackwell (10.1111); 2012. pp. 1–18. pmid:22462663
  41. 41. Palmer KJ, Hughes H, Stephens DJ. Specificity of cytoplasmic dynein subunits in discrete membrane-trafficking steps. Holzbaur E, editor. Mol Biol Cell. 2009;20: 2885–2899. pmid:19386764
  42. 42. Myers PZ. Spinal motoneurons of the larval zebrafish. J Comp Neurol. Wiley-Blackwell; 1985;236: 555–561. pmid:4056102
  43. 43. Tanaka E, Sabry J. Making the connection: cytoskeletal rearrangements during growth cone guidance. Cell. 1995;83: 171–176. pmid:7585934
  44. 44. Brandt R. Cytoskeletal mechanisms of axon outgrowth and pathfinding. Cell Tissue Res. 1998;292: 181–189. pmid:9560461
  45. 45. Blanquie O, Bradke F. Cytoskeleton dynamics in axon regeneration. Curr Opin Neurobiol. 2018;51: 60–69. pmid:29544200
  46. 46. Ruschel J, Hellal F, Flynn KC, Dupraz S, Elliott DA, Tedeschi A, et al. Axonal regeneration. Systemic administration of epothilone B promotes axon regeneration after spinal cord injury. Science. American Association for the Advancement of Science; 2015;348: 347–352. pmid:25765066
  47. 47. Hellal F, Hurtado A, Ruschel J, Flynn KC, Laskowski CJ, Umlauf M, et al. Microtubule stabilization reduces scarring and causes axon regeneration after spinal cord injury. Science. American Association for the Advancement of Science; 2011;331: 928–931. pmid:21273450
  48. 48. Sengottuvel V, Leibinger M, Pfreimer M, Andreadaki A, Fischer D. Taxol facilitates axon regeneration in the mature CNS. J Neurosci. Society for Neuroscience; 2011;31: 2688–2699. pmid:21325537
  49. 49. Grabham PW, Seale GE, Bennecib M, Goldberg DJ, Vallee RB. Cytoplasmic Dynein and LIS1 Are Required for Microtubule Advance during Growth Cone Remodeling and Fast Axonal Outgrowth. Journal of Neuroscience. 2007;27: 5823–5834. pmid:17522326
  50. 50. Adames NR, Cooper JA. Microtubule interactions with the cell cortex causing nuclear movements in Saccharomyces cerevisiae. The Journal of Cell Biology. 2000;149: 863–874. pmid:10811827
  51. 51. Ligon LA, Tokito M, Finklestein JM, Grossman FE, Holzbaur ELF. A direct interaction between cytoplasmic dynein and kinesin I may coordinate motor activity. J Biol Chem. American Society for Biochemistry and Molecular Biology; 2004;279: 19201–19208. pmid:14985359