Axons in the CNS do not regrow after injury, whereas lesioned axons in the peripheral nervous system (PNS) regenerate. Lesioned CNS axons form characteristic swellings at their tips known as retraction bulbs, which are the nongrowing counterparts of growth cones. Although much progress has been made in identifying intracellular and molecular mechanisms that regulate growth cone locomotion and axonal elongation, a comprehensive understanding of how retraction bulbs form and why they are unable to grow is still elusive. Here we report the analysis of the morphological and intracellular responses of injured axons in the CNS compared with those in the PNS. We show that retraction bulbs of injured CNS axons increase in size over time, whereas growth cones of injured PNS axons remain constant. Retraction bulbs contain a disorganized microtubule network, whereas growth cones possess the typical bundling of microtubules. Using in vivo imaging, we find that pharmacological disruption of microtubules in growth cones transforms them into retraction bulb-like structures whose growth is inhibited. Correspondingly, microtubule destabilization of sensory neurons in cell culture induces retraction bulb formation. Conversely, microtubule stabilization prevents the formation of retraction bulbs and decreases axonal degeneration in vivo. Finally, microtubule stabilization enhances the growth capacity of CNS neurons cultured on myelin. Thus, the stability and organization of microtubules define the fate of lesioned axonal stumps to become either advancing growth cones or nongrowing retraction bulbs. Our data pinpoint microtubules as a key regulatory target for axonal regeneration.
- retraction bulb
- dorsal root ganglia neurons (DRG neurons)
- in vivo imaging
- growth cone
Growth cones regulate neurite extension and sense guidance cues during development (Huber et al., 2003). After injury, the formation of a growth cone at the tip of a transected axon is a crucial step during subsequent axonal regeneration. The tips of lesioned axonal stumps located in the PNS transform into growth cones of stereotypic spiky morphology capable of sustained growth. On the contrary, lesioned CNS axons form swellings termed “retraction bulbs” at the tip of their proximal stumps, which are oval structures and lack a regenerative response (Ramon y Cajal, 1928; Li and Raisman, 1995; Hill et al., 2001). Although there is a growing interest in identifying the intrinsic mechanisms that prevent axonal regeneration in these end structures (Silver and Miller, 2004; Tom et al., 2004; Steinmetz et al., 2005), the current knowledge about retraction bulbs remains fragmentary.
Growth cones contain the machinery for movement and axonal extension consisting of a complex interplay of different intracellular events. For example, mitochondria concentrate in the tip of the growing axon to provide energy necessary for axon formation (Morris and Hollenbeck, 1993; Bradke and Dotti, 1997). Axon growth also depends on continuous membrane supply from the soma to support the surface expansion of the growing axon (Craig et al., 1995; Dai and Sheetz, 1995; Vogt et al., 1996). Notably, microtubules and their dynamic rearrangements are essential for axon outgrowth (Forscher and Smith, 1988; Sabry et al., 1991; Tanaka and Kirschner, 1991; Dent and Gertler, 2003).
Our knowledge about the intrinsic mechanisms that restrain axons from regrowing is mostly based on in vitro models, including the growth cone collapse and the stripe assays. Although such assays helped discover important repulsion signals, including Semaphorins and Ephrins (Raper and Kapfhammer, 1990; Drescher et al., 1995), they have limitations in recapitulating the morphology and intracellular mechanisms of stalled axons in vivo, e.g., after spinal cord injury. For example, in the growth cone collapse assay, growth cones retract their filopodia and lamellipodia, showing enhanced retrograde transport of varicosities, resulting in rapid thinning of the fan-like growth cone after ligand stimulation (Fan et al., 1993), which is distinct from the bulbous structure of a retraction bulb. Recently, a cell culture model that produces voluminous axonal tips reminiscent to retraction bulbs has been established using a proteoglycan–laminin gradient (Tom et al., 2004; Steinmetz et al., 2005).
However, the underlying mechanisms of retraction bulb formation and axon stalling still remain unclear. To investigate processes that malfunction in injured CNS axons and generate an obstacle for regeneration, we analyzed the intracellular structure of retraction bulbs compared with growth cones. We chose to work with dorsal root ganglia (DRG) neurons because their peripheral axonal branches form growth cones after lesioning and regenerate, whereas the central branches form retraction bulbs and do not regenerate.
Materials and Methods
We used green fluorescent protein-M (GFP-M) and yellow fluorescent protein-H (YFP-H) transgenic mice (8–12 weeks of age; 15–30 g) expressing GFP/YFP under the control of the neuron-specific Thy-1 promoter (Feng et al., 2000). All animal experiments were performed in accordance with the animal handling laws of the government (Regierung von Oberbayern, Number 209.1/211-2531-115/02).
Dorsal column lesion and postoperative care.
The mice were anesthetized with a mixture of midazolam (Dormicum; 2 mg/kg), medetomidine (Domitor; 0.15 mg/kg), and fentanyl (0.05 mg/kg) injected intraperitoneally. The lamina at the level of T8/9 was removed by laminectomy to expose the dorsal columns at the thoracic level. Fine iridectomy scissors (FST, Heidelberg, Germany) were used to transect the dorsal columns bilaterally. The lamina was closed, and the skin was stapled. Two to three hours after surgery, animals were woken up by subcutaneous injection of a mixture of flumazenil (Antisedan; 0.2 mg/kg), atipamezole (Anexate; 0.75 mg/kg), and naloxone (Narcanti; 0.12 mg/kg). The animals were kept on a heating pad for the following 24 h. Postsurgical animal care was done as follows: the bladders were expressed every day twice; antibiotic (10 μl of 7.5% Borgal solution; Hoechst Russel Vet, Frankfurt, Germany) was given subcutaneously once daily, and buprenorphine (0.15 mg/kg) was given once every 12 h after surgery, in total three times.
Sciatic nerve lesion.
The mice were anesthetized as described above. The sciatic nerve was exposed ∼2.5 cm from the DRG cell bodies by a small incision and crushed with forceps for 10 s. The incision was sutured, and the skin was stapled. The anesthesia was reversed, and the animals were transferred to a heating pad for the following 24 h.
Perfusion and sectioning.
Animals were killed and perfused at the defined time points. In brief, animals were anesthetized with 5% chloral hydrate solution (prepared in saline) and perfused intracardially with 0.1 m PBS at a speed of 3 ml/min for 5 min, immediately followed by 4% paraformaldehyde (PFA) in PBS for 20 min for immunostainings and for 45 min for morphology analysis. The spinal cord or sciatic nerve lesion sites were carefully dissected and postfixed in 4% PFA (at room temperature for 30 min for immunostainings, at 4°C overnight for morphology analysis). The tissues were incubated in 15% sucrose at room temperature for 4 h and in 30% sucrose overnight. The tissues were embedded in optimal cutting temperature compound (Tissue-Tek; Miles, Elkhart, IN) and sectioned longitudinally by a cryostat (CM 3050; Leica, Wetzlar, Germany) at 10 μm. For size quantifications, we made either thick cryostat sections (40 μm) or whole mounts for confocal observation.
The primary antibodies used were anti-glutamated (Glu)-tubulin (rabbit) at 1:500 dilution (Millipore, Billerica, MA), anti-Tyr-tubulin (rat) at 1:500 (Abcam, Cambridge, MA), and anti-golgin-160 (rabbit) at 1:200 (a gift from Dr. Francis Barr, Max Planck Institute of Biochemistry, Martinsreid, Germany). Tissue sections were washed with PBS twice for 10 min and then incubated with blocking solution containing 10% goat serum and 0.3–0.5% Triton X-100 in PBS at room temperature for 1 h, washed with PBS, and incubated with primary antibody (in blocking solution with 5% goat serum) at 4°C overnight. The sections were washed with PBS and incubated with an anti-rabbit secondary antibody Alexa 568 (Invitrogen, Eugene, OR). The sections were mounted with water-based mounting medium (Polysciences, Warrington, PA).
DRG neuron culture and nocodazole application.
Dorsal root ganglia were isolated from L4 and L5 spinal levels of adult female Sprague Dawley rats (200–250 g). Ganglia were enzymatically digested in 1% collagenase type I (Worthington Biochemical, Lakewood, NJ) for 90 min and then in 0.25% trypsin (Sigma-Aldrich, St. Louis, MO) at 37°C for 15 min. Enzymatic digestion was stopped by addition of Neurobasal medium (Invitrogen) containing 5% horse serum, and neurons were recovered by centrifugation at 500 rpm for 5 min. Dissociated neurons were cultured on laminin-coated dishes (50 μg/ml; Roche, Indianapolis, IN) in Neurobasal media complemented with B-27 supplement, l-glutamine, and penicillin-streptomycin. Cells were incubated at 37°C in a humidified atmosphere containing 5% CO2. After 48 h, cells were fixed with 4% paraformaldehyde and stained for βIII-tubulin (TuJ1; Covance, Princeton, NJ) and Glu-tubulin antibodies.
Cerebellar granule neurons culture and taxol application.
Granule neurons were isolated from the cerebellar cortex of 9-d-old rats and cultured as described previously (Hatten, 1985). Briefly, the cerebellar cortex was trypsinized, followed by trituration in a Trypsin/DNase mix (10 and 1 mg/ml, respectively) in Tyrode solution (CMF-PBS). The granule cells were suspended in Neurobasal medium (Invitrogen) complemented with B-27 supplement (Invitrogen), 10% calf serum (Invitrogen), glutamine (Invitrogen), and 1% penicillin-streptomycin (Invitrogen) and purified by preplating the cell suspension on poly-d-lysine (Sigma-Aldrich)-coated dishes. The cells were plated at 2.5 × 106 per well in eight-well chamber slides [Lab-Tek (Campbell, CA) system; Nunc (Pittsburgh, PA)] coated with 1 mg/ml poly-d-lysine alone or together with myelin as outgrowth-inhibitory substrate. The myelin substrate was prepared as described previously (Neumann et al., 2002), and 1 μg of protein was dried on the poly-d-lysine-precoated dishes per well overnight.
Eighteen hours after plating, cells were fixed with 4% paraformaldehyde and immunostained with the βIII-tubulin neuronal marker (TuJ1; Covance). The longest neurite per neuron was measured in duplicates on an average of 500 cells per condition, and the results were expressed as the percentage of neurites exhibiting >60 μm length. Each experiment was repeated at least three times.
We acquired confocal images with a Leica SP2 confocal microscope system in sequential scanning mode.
Animals were perfused as described above by using Lewis Shute fixative (Lewis and Shute, 1969). The tissue was dissected without drying and postfixed in Lewis Shute fixative and then in osmium tetroxide. Afterward, the tissue was dehydrated and embedded in Araldite, cut by an ultramicrotome (LKB, Mt. Waverley, Australia) at 50 nm and poststained with lead citrate and uranyl acetate by an ultrastainer (LKB). Electron microscopy (EM) images of the sections were acquired using a Zeiss (Thornwood, NY) EM 10 electron microscope.
In vivo imaging.
We used an Olympus (Tokyo, Japan) SZX-12 fluorescent stereomicroscope equipped with 1× Plan Apochromat objective and 1.6× Plan Fluorite objective (Olympus). A ColorView II camera integrated to the microscope was used to capture images through Analysis FIVE software (Soft Imaging System, Münster, Germany). Animals were anesthetized as described above.
Treatment of peripheral axons in vivo.
Sciatic nerve lesion was performed as described above. After 24 h, animals were reanesthetized, and 10 μl of 330 μm nocodazole (diluted in 5% DMSO in PBS; Sigma-Aldrich) or 5% DMSO alone was applied onto the lesion site with a pipette. The injury site was closed, and animals were perfused 24 h after nocodazole or DMSO application for staining and imaging. For assessing peripheral axon growth, the animals were reanesthetized 1–2 d after lesioning, and nocodazole, taxol (10 μl of 1 μm), or DMSO was applied in the same manner. The animals were either observed for 6 h and then killed or reimaged 24 h later, depending on the experimental protocol.
Treatment of central axons in vivo.
After removing the laminas at T11/12 level, a small unilateral lesion was applied with vannas spring scissors (FST) by transecting only the superficial axons of the dorsal columns. We imaged the injury site before and after lesion by acquiring serial images. The animals were kept on a heating pad during the imaging session. Ten microliters of 1 μm taxol (diluted in saline; Sigma-Aldrich) or saline as control was applied onto the lesion site each hour, starting immediately after lesioning. The lesion sites were visualized by in vivo imaging as described above. The animals' anesthesia was boosted after 3 h. At the end of the observation period, animals were perfused and prepared for the confocal imaging as described above.
Size, distance, and length quantifications.
The size of retraction bulbs and growth cones was quantified as follows: the maximum diameter of the retraction bulb and growth cone at the tip of the axon was measured and divided by the diameter of the cylindrical shaft of the axon.
The retraction distance of taxol- and saline-treated axons to the lesion site was quantified as follows: the lesion site was outlined from the in vivo binocular images. We then measured the distance of the axonal tips to the proximal edge of the lesion.
To measure neurite length in dorsal root ganglia neurons, we took pictures from each well (80 cells per well on average) using a confocal microscope and a 10× objective. We then analyzed the data using Amira 4.1 (Mercury Computer Systems, Chelmsford, MA). The mean length has been calculated by measuring the total number of pixels in the images from which the pixels of the cell bodies have been subtracted and dividing by the average pixel number per micrometer of the axons and the total number of cell bodies in the image [total length per neuron = total number of pixels (only axons) × (pixel number per micrometer)−1 × (number of neurons)−1].
Image processing and data evaluation.
Angle deviations, numbers of vesicles and mitochondria, size of end structures, and distance to the lesion site were quantified with Analysis FIVE software. The images were assembled with Photoshop (Adobe Systems, San Jose, CA) and Canvas (ACD Systems, Miami, Florida). The diagrams were drawn with Illustrator (Adobe Systems). For the three-dimensional (3D) reconstructions, the quantification of vesicle volumes and the total length of DRG neurons after in vitro nocodazole treatment were measured using Amira 4.1. The statistical analysis was performed with Excel (Microsoft, Seattle, WA), and statistical significance (p < 0.05) was calculated using the two-tailed, unpaired t test.
Morphological changes of growth cones and retraction bulbs
The peripheral axonal branches of the DRG neurons run in the PNS, where they generate growth cones and regrow after nerve injury (Fig. 1 A). In contrast, the central axonal branches coursing in the dorsal columns of the spinal cord do not regrow after damage, and the lesioned proximal axonal terminals form retraction bulbs (Fig. 1 A). To analyze the differences between retraction bulbs and growth cones in vivo, we used transgenic mice expressing GFP in a small subset of neurons, allowing a clear visualization of individual axons (Fig. 1 B–K; supplemental Fig. S1, available at www.jneurosci.org as supplemental material) (Feng et al., 2000; Kerschensteiner et al., 2005). We induced growth cone formation by lesioning the sciatic nerve, which contains the peripheral axonal branches from the lumbar 4 and 5 DRG neurons (Fig. 1 A–E; supplemental Fig. S1C–E, available at www.jneurosci.org as supplemental material). Retraction bulb formation, on the other hand, was induced by lesioning the dorsal columns at the thoracic 8 and 9 level (Fig. 1 A,F–K; supplemental Fig. S1F–H, available at www.jneurosci.org as supplemental material).
Peripherally axotomized DRG neurons in vivo generated a slim growth cone that remained morphologically constant over time (Fig. 1 B–E). Although growth cones of DRG neurons in cell culture have filopodia (supplemental Fig. S2, available at www.jneurosci.org as supplemental material), they rarely show such structures in vivo. The maximal width of the in vivo growth cones was comparable with the diameter of the axonal shaft: the growth cone (tip)/shaft ratio was 1.31 ± 0.14 at day 1 (Fig. 1 B,C) (n = 113 growth cones), 1.51 ± 0.15 at day 4 (n = 48 growth cones), and 1.20 ± 0.20 at 1 week after sciatic lesion (Fig. 1 D,E) [n = 21 growth cones; average ± SD; n ≥ 7 mice for each time point (Fig. 1 L)]. Later time points were difficult to visualize because of the high velocity of axonal regeneration in the PNS.
After axotomizing the central axonal branches, retraction bulbs formed at the proximal tips. These bulbs were typically round or oval with a much larger diameter than their axons and lacked any kind of extensions (Fig. 1 F–K). Retraction bulbs increased approximately three times in size from 1 d to 5 weeks after injury (Fig. 1 F–K,L). The retraction bulb (tip)/axonal shaft ratio was 4.09 ± 0.73 at 1 d (Fig. 1 F,G) (n = 109 retraction bulbs), 8.49 ± 0.51 at 1 week (Fig. 1 H,I) (n = 111 retraction bulbs), and 11.79 ± 0.49 at 5 weeks after injury (Fig. 1 J,K) [n = 92 retraction bulbs; average ± SD; n ≥ 8 mice for each time point (Fig. 1 L)]. In contrast to the slim growth cones that were continuous with the peripheral axonal shafts, the surface area of bulbous structures increased over time (Fig. 1 M). Some of the axons formed smaller bulbs on their shafts in addition to the bigger terminal bulbs [supplemental Fig. S1G (available at www.jneurosci.org as supplemental material), arrow; Fig. 1 I,K]. We next aimed to address the underlying intracellular differences that could cause the formation of retraction bulbs and stalling of axons.
Retraction bulbs contain mitochondria and trans-Golgi-network-derived vesicles
Sustained axonal growth requires various intracellular activities (e.g., transport of vesicles and cytoskeletal dynamics) that need a high energy supply from mitochondria (Morris and Hollenbeck, 1993). Using electron microscopy, we analyzed the mitochondrial morphology and density in growth cones and retraction bulbs at various time points. The mitochondria of the retraction bulbs (Fig. 2 C,D) were indistinguishable from the mitochondria of growth cones (Fig. 2 A,B) and did not show characteristics of degenerating mitochondria that typically contain swollen membranes and loss of cristae (Kong and Xu, 1998). Mitochondrial density in retraction bulbs at 4 d, 2 weeks, and 5 weeks after injury (3.63 ± 0.61, 3.9 ± 0.62, and 4.75 ± 0.66 mitochondria/μm2, respectively; average ± SEM; n ≥ 7 retraction bulbs per time point) was significantly higher (p < 0.05) (Fig. 2 M) than in growth cones (1.76 ± 0.47; average ± SEM; n = 8 growth cones).
To investigate whether lack of membrane traffic underlies both retraction bulb formation and axonal stalling after CNS injury we assessed post-Golgi trafficking by coimmunostaining GFP-labeled growth cones and retraction bulbs using the trans-Golgi-derived vesicle marker golgin-160 (Misumi et al., 1997). Post-Golgi vesicles accumulated both in the growth cones (Fig. 2 E–H) and retraction bulbs (Fig. 2 I–L). Similar results were obtained by electron microscopy (Fig. 2 A–D, arrows). The density of smooth, small-sized vesicles was significantly higher in the retraction bulbs at 4 d, 2 weeks, and 5 weeks after injury than in the growth cones (p < 0.05) (Fig. 2 N) (24.6 ± 3.86, 35.7 ± 6.54, and 51.17 ± 6.52 vesicles/μm2, respectively, for retraction bulbs; average ± SEM; growth cones, 12.15 ± 2.5; n ≥ 7 retraction bulbs per time point and n = 8 growth cones). Additionally, vesicle density in the retraction bulbs significantly increased over time (p < 0.01) (Fig. 2 N, from 4 d to 5 weeks after injury). Thus, it appears that vesicles are accumulating over time in the retraction bulbs. We quantified the vesicle densities in the end structures also on 3D images obtained after reconstruction of confocal stacks and found results similar to the quantifications on the EM images (Fig. 2 H,L,O) [26.6 ± 1.3% vesicles/volume (vol) in retraction bulbs at 2 weeks and 14.1 ± 3.7% vesicles/vol in growth cones; average ± SD of the total volume of the end structure; n = 4 3D images per condition]. Altogether, we could not find any apparent deficiencies in energy supply or membrane trafficking that hamper axon extension after spinal cord injury.
Retraction bulbs contain disorganized microtubules
Microtubules play a key role in axonal growth and guidance (Yamada et al., 1970; Forscher and Smith, 1988; Suter et al., 1998; Dent et al., 1999; Dent and Gertler, 2003). They form the backbone of the axonal shafts and core domain of growth cones, giving stability to those structures and enabling organelle transport (Hirokawa and Takemura, 2005). In addition, the dynamic microtubules protrude through the peripheral regions of growth cones, enabling axon elongation (Kabir et al., 2001). We therefore investigated whether the microtubular array differs in nongrowing retraction bulbs and regenerating growth cones. To assess the integrity of microtubules, we coimmunostained GFP-labeled axonal shafts, growth cones, and retraction bulbs for detyrosinated tubulin using an anti-Glu-tubulin antibody and tyrosinated tubulin using an anti-Tyr antibody as markers for stable and dynamic microtubules, respectively (Westermann and Weber, 2003). We found that detyrosinated (stable) microtubules were tightly bundled and parallel aligned in the growth cones and their axonal shaft (Fig. 3 A–D) (n = 21 growth cones). The parallel microtubule arrays were observed all over the growth cone (Fig. 3 D, arrowheads). Tyrosinated (dynamic) microtubules also showed a bundling that reaches to the tips of the growth cones (supplemental Fig. S3E–H, available at www.jneurosci.org as supplemental material) (n = 12 growth cones), similar to what has been demonstrated in cultured neurons (Schaefer et al., 2002; Dent and Gertler, 2003). In contrast, stable microtubules were highly disorganized in retraction bulbs (Fig. 3 E–H) (n = 19 retraction bulbs). Although some regions contained accumulation of stable microtubules (Fig. 3 F, white arrow), other regions were devoid of microtubule filaments (Fig. 3 F, white arrowhead). The orientation of different microtubules was highly heterogeneous (Fig. 3 F, yellow arrows). For instance, none of the microtubules that are indicated with yellow arrowheads in the 3D reconstruction (Fig. 3 H) were parallel to each other. Interestingly, dynamic microtubules were also found in retraction bulbs, albeit seemingly to a smaller extent than in the growth cones (supplemental Fig. S3A–D, available at www.jneurosci.org as supplemental material) (n = 7 retraction bulbs), suggesting that the microtubules are still dynamic and can integrate new tubulin dimers. However, these dynamic microtubules in retraction bulbs were disordered (Fig. S3C, yellow arrows). Moreover, dynamic microtubules were almost absent from the tips of retraction bulbs (Fig. S3C, white arrowhead) but accumulated in the more central domain (Fig. S3C, white arrow). These results suggest that the mislocalization of dynamic microtubules in the nongrowing bulbs could account for axon stalling.
We further quantified the deviation of microtubules from the axonal axis by analyzing electron micrographs of axonal terminals. In unlesioned controls, the microtubules aligned in parallel to the axonal axis: 99% of the microtubules deviated <30° from the axonal axis [a representative image is shown in Fig. 3 I; the microtubules of the same image were traced with black lines for clearer visualization in Fig. 3 J; a higher magnification of the marked area, showing the angle of deviation, is presented in Fig. 3 K (n = 7 axonal shafts)]. In growth cones, 97.0% of the microtubules deviated <30° from the axonal axis, similar to the unlesioned controls (Fig. 3 L–N) (n = 8 growth cones). In retraction bulbs, only 51% of the microtubules deviated <30° from the axonal axis; instead, 35% of the microtubules deviated between 30 and 60°, and 13% deviated >60° from the axonal axis (Fig. 3 O–R) (n = 9 retraction bulbs). These results show that microtubules in retraction bulbs are disorganized and dispersed compared with unlesioned axons and growth cones (p < 0.001) (the quantification of the data are shown in Fig. 4).
Microtubule destabilization converts a growth cone into a retraction bulb-like structure and halts axonal growth in vivo
Our observation of disorganized microtubules in retraction bulbs led us to examine whether destabilization of microtubules in a growth cone would be sufficient to transform it into a retraction bulb. To address this point, we injured the sciatic nerve of GFP-M mice to obtain regenerating growth cones, as before, and applied 10 μl of the microtubule-disrupting drug nocodazole (330 μm) to the site of the lesioned sciatic nerve 1 d later. During examination of the sciatic nerve 24 h after nocodazole treatment, we found that 42.3 ± 13.2% of the axons (average ± SD; n = 165 axons from 11 mice) had transformed their growth cones into a bulb (Fig. 5 D–F). Application of DMSO, which served as a negative control, did not cause bulb formation (Fig. 5 A–C). The tip/axonal shaft ratio of nocodazole-induced bulbs (4.25 ± 0.56; average ± SD; n = 70 from 12 mice) was significantly higher than in both lesioned, untreated (1.45 ± 0.13; average ± SD; n = 53 from 8 mice) and lesioned, DMSO-treated (1.64 ± 0.46; average ± SD; n = 62 from 12 mice) controls (Fig. 5 G) (p ≤ 0.001 for both). These nocodazole-induced peripheral bulbs showed a size increase similar to the retraction bulbs seen in the CNS 1 d after spinal cord injury (4.09 ± 0.73) (Fig. 1 F,G,L). When we analyzed the nocodazole-induced peripheral bulbs, we found a dispersed microtubule organization, similar to that in retraction bulbs of the CNS (compare Figs. 5 H–K, 3 E–H). Lower concentrations of nocodazole did not cause bulb formation (10 μl of 3.3 or 33 μm nocodazole) (supplemental Fig. S4A–J, available at www.jneurosci.org as supplemental material).
To assess the growth capacity of the nocodazole-induced peripheral bulbs, we performed in vivo imaging and followed the advance of injured peripheral axonal endings (Fig. 5 L–O). Two days after sciatic nerve injury, the lesion sites containing the growth cones were treated with either nocodazole or vehicle (DMSO). We imaged axons at the time of treatment and 6 h later and measured their extension. Nocodazole treatment (10 μl of 330 μm) significantly (p < 0.05) inhibited the growth of axons compared with control treatment [Fig. 5 L,M, green arrowheads (control); Fig. 5 N,O, red arrowheads (nocodazole)]. All control animals showed axonal growth of 20–100 μm within the observation period. In contrast, nocodazole-treated animals showed very little axonal regrowth: only two of seven animals extended their axons up to 10 μm, whereas the other nocodazole-treated animals did not elongate their axons during the observation period (Fig. 5 P). Lower concentrations of nocodazole did not show any growth-inhibitory effect (10 μl of 3.3 or 33 μm nocodazole) (supplemental Fig. S4K–P, available at www.jneurosci.org as supplemental material).
Together, our results demonstrate that destabilization of microtubules in vivo is sufficient to generate retraction bulb-like structures from a growth cone, as judged by morphological, cytoskeletal, and growth criteria.
Microtubule destabilization induces the formation of bulbous axon terminals and restricts axon outgrowth in cell culture
To address whether nocodazole-induced axonal stalling and morphological changes of the growth cones function cell autonomously, we attempted to generate retraction bulb-like structures in isolated DRG neurons in cell culture. DRG neurons were dissected and cultured as described previously (Neumann et al., 2002). One day after plating, when DRG neurons formed several axons with growth cones at their tips, they were treated with various concentrations of nocodazole for another day. Whereas only a minor fraction of control neurons had tips with a bulbous, club-like morphology, nocodazole-treated neurons had significantly more bulbous endings (Fig. 6 A–J) (9.6 ± 2.3% in controls, 29.1 ± 4.4% in 75 nm nocodazole, 41.1 ± 8.2% in 250 nm nocodazole; p < 0.001; average ± SD; data derived from three independent cultures per condition). Similar to the retraction bulbs formed after spinal cord injury, nocodazole-induced bulbous endings possessed disorganized stable and dynamic microtubules compared with the growth cones (Fig. 6 K,L; supplemental Fig. S5, available at www.jneurosci.org as supplemental material).
The growth of the nocodazole-treated axons was significantly (p < 0.01) inhibited compared with controls. The total length of the axons per DRG neuron 2 d in culture (treated from 24 to 48 h) was 2849 ± 46 μm in controls but reduced to 2132 ± 138 μm in 75 nm nocodazole and 1555 ± 142 μm in 250 nm nocodazole (Fig. 6 M–P) (average ± SEM; data derived from three independent cultures per condition). After nocodazole treatments, the total number of cells did not change (data not shown). Thus, destabilization of microtubules is sufficient to cause formation of bulbous endings reminiscent of retraction bulb-like structures by their morphology, microtubule structure, and growth ability.
Microtubule stabilization interferes with the formation of retraction bulbs
Because microtubule destabilization appears to be crucial element in the generation of retraction bulbs, we wondered whether microtubule stabilization would be sufficient to prevent the formation of retraction bulbs after CNS injury. To this end, we applied taxol, a microtubule-stabilizing drug, to the injured spinal cord and assessed the alteration of the axonal stumps' terminals by in vivo imaging. We performed a small unilateral lesion in the dorsal columns of GFP-M mice at the T12 level (Fig. 7 A,B), and the injury sites were bathed in either the microtubule-stabilizing drug taxol or in vehicle (saline). We then continuously observed the behavior of the axons during the subsequent 6 h, using a binocular microscope (Kerschensteiner et al., 2005; Misgeld and Kerschensteiner, 2006) (Fig. 7 C,D). In control conditions, we started to observe formation of apparent retraction bulbs 1 h after injury (Fig. 7 D). At 6 h after injury, already 71.3 ± 9.1% of the axons (average ± SD; n = 18 mice) had formed retraction bulbs at their tips (Fig. 7 D,F,M). However, when 1 μm taxol was repetitively applied (10 μl/h), starting immediately after the injury, only 22.8 ± 13.1% (average ± SD; n = 11 mice) of the axons developed bulbs (defined by a tip/axon ratio of >2) in the first 6 h (Fig. 7 C,E,M). In addition, the size of the axon terminals formed after taxol treatment was smaller (Fig. 7 G–I) [tip/axon ratio, 2.53 ± 0.54 (average ± SD) for the five biggest retraction bulbs per animal] than the retraction bulbs of control treatment (Fig. 7 J–L) [4.14 ± 0.44 (average ± SD); n = 11 mice for each condition; p < 0.001]. The microtubules of taxol-treated axonal endings were bundled and aligned in parallel to the axonal axis resembling the cytoskeletal structure of in vivo growth cones (compare Figs. 7 N–R, 3 A–D). Interestingly, some axons of taxol-treated animals also showed decreased acute axonal degeneration (Kerschensteiner et al., 2005) and Wallerian degeneration as shown by the reduced axonal retraction from the lesion site. In six of 11 taxol-treated animals, the proximal axonal stumps were as close as 25 μm to the injury site 6 h after injury (Fig. 7 E, red arrow), whereas only one of 11 controls showed such minimal retraction (Fig. 7, compare E, F). These results suggest that stabilization of microtubules prevents the formation of retraction bulbs and interferes with axonal retraction after CNS lesioning. To test whether the effect of taxol is specific for inhibiting degeneration of central axonal branch or whether it can change the growth of the peripheral branch after injury, we applied the same concentration of taxol onto the regenerating axons after a sciatic nerve lesion. We observed that 1 μm taxol application did not significantly alter the growth speed of the axons compared with controls. The average growth of the axons was 572 ± 277 μm in the taxol-treated group (supplemental Fig. S6A,B,E, available at www.jneurosci.org as supplemental material) and 511 ± 276 μm in the DMSO-treated group (supplemental Fig. S6C,D,E, available at www.jneurosci.org as supplemental material) within 24 h after treatment (average ± SD; n ≥ 7 mice per group). This suggests that taxol at the concentration used interferes with the growth restraint inflicted on axons but does not change the general growth rates of axons.
Microtubule stabilization enhances the growth capacity of neurons cultured on myelin
We further investigated whether taxol application could enhance axonal growth of neurons cultured on an inhibitory substrate. Cultured DRG neurons contain axons that resemble peripheral axonal branches according to their growth rates (Smith and Skene, 1997). We therefore decided to use CNS neurons to study growth of axons on inhibitory substrates. Dissociated cerebellar granule neurons (CGNs) were plated on poly-lysine or CNS myelin (Niederost et al., 1999) and treated with taxol or vehicle. When granule neurons were cultured on poly-lysine, 42.9 ± 4.1% of neurons formed long neurites within 18 h (Fig. 8 A,B,I). In contrast, myelin inhibited the formation of long neurites: only 7.4 ± 2.6% of neurons were able to form a process longer than 60 μm (Fig. 8 E,F,I). Remarkably, in the presence of taxol (10 nm), a significant amount of neurons plated on myelin showed enhanced axonal growth; 29.9 ± 2.35% of the cells contained neurites longer than 60 μm (p < 0.001) (Fig. 8 G–I). Notably, taxol application did not change the growth capacity of CGNs when they were plated on poly-lysine: 40.4 ± 3.3% of neurons had processes longer than 60 μm (Fig. 8 C,D,I) (average ± SD; n ≥ 3 independent experiments per condition). These results show that stabilization of microtubules in culture increases the growth ability of neurons in the presence of an inhibitory environment.
Retraction bulbs are the morphological hallmarks of lesioned axons that do not regenerate (Ramon y Cajal, 1928). We have shown that one of the main features of retraction bulb formation is the loss of the microtubule organization (Fig. 9). Disorganizing the microtubules of growth cones using nocodazole was sufficient to generate retraction bulbs as well as to inhibit axonal growth both in vitro and in vivo. Conversely, preventing microtubule disorganization by the microtubule-stabilizing drug taxol blocks the formation of retraction bulbs. Moreover, we demonstrate that microtubule stabilization is sufficient to induce elongation of CNS axons cultured on a growth-inhibitory substrate.
The basic intracellular events of retraction bulb formation and maintenance
Formation of axonal swellings is one of the early manifestations of axonal degeneration. In our experiments, the first retraction bulbs start to form within 1 h, consistent with previous observations (Kerschensteiner et al., 2005), and they possess dynamic morphologies during initial hours of injury. Subsequently, they enlarge in size over weeks.
In aiming to identify the mechanisms underlying retraction bulb formation and lack of axonal growth, we assessed effects on membrane trafficking, energy supply, and microtubules. Our results suggest that trans-Golgi-derived vesicles and mitochondria reach and accumulate in the retraction bulbs, indicating that they are not restricting factors of axon regeneration after CNS injury. Indeed, ongoing membrane trafficking of stalled axons might be one cause for the expansion of the retraction bulbs over time, together with endocytosis (Tom et al., 2004). Whereas growth cones in vivo have bundled and parallel-aligned stable and dynamic microtubules that reach the tip, we found that retraction bulbs contain disorganized and dispersed stable and dynamic microtubules. Notably, dynamic microtubules are mislocalized in retraction bulbs. This contrasts with in vivo growth cones, which contain bundled dynamic microtubules that extend to the axonal tips. Reorganization of dynamic microtubules is crucial to generate a pushing force against the plasma membrane (Dogterom et al., 2005). Dynamic microtubules probe the actin-rich peripheral regions of in vitro growth cones and continuously project into lamellipodia and filopodia, showing the typical dynamic behavior of microtubules necessary for axon extension (Kabir et al., 2001; Schaefer et al., 2002). Thus, it appears that disorganized microtubules present in retraction bulbs are not able to support axon extension.
Interestingly, slowly growing or pausing growth cones in vitro share some similarities with retraction bulbs. For example, pausing growth cones expand their surface area and show rearrangements of their microtubules (Sabry et al., 1991; Tanaka and Kirschner, 1991; Dent et al., 1999). Specifically, WNT-7a causes axon inhibition and the generation of enlarged mossy fiber growth cones that possess unbundled microtubules reminiscent to the microtubules in retraction bulbs (Hall et al., 2000).
Manipulating the microtubule network to convert the end structures
Modification of microtubules with pharmacological drugs can change the direction (Buck and Zheng, 2002), morphology, and elongation of the axons (Pan et al., 2003). Our findings demonstrate that microtubule destabilization using nocodazole is sufficient to produce retraction bulb-like structures from growth cones and to block axonal growth both in culture and in vivo after peripheral injury.
We also demonstrate that stabilization of microtubules using taxol after CNS injury inhibits dispersion of the microtubules, preventing retraction bulb formation and axonal degeneration. One way to interpret these results is that taxol could inhibit microtubule catastrophe events that occur after CNS injury, probably caused by a rapid intra-axonal accumulation of calcium (Balentine and Spector, 1977; O'Brien et al., 1997). Interestingly, pharmacological inhibition of calpains, a group of proteases that cleaves the major proteins, including cytoskeletal proteins (George et al., 1995), also has a neuroprotective effect in the optic nerve (Araujo Couto et al., 2004) and in DRG neurons (Kerschensteiner et al., 2005).
In addition, we show that taxol treatment increases the growth ability of cultured postnatal CGNs when plated on CNS myelin, which is an established cell culture assay that resembles some aspects of the inhibitory environment of the injured spinal cord. Interestingly, taxol-treated neurons do not show enhanced growth when plated on poly-lysine, suggesting that taxol enables axons to overcome growth restraint rather than enhancing their general growth potential. Consistent with this interpretation, we demonstrate that taxol-treated injured peripheral axons of DRG neurons do not change their growth rate.
Together, our in vitro and in vivo findings present promising insights to study the therapeutic potentials of microtubule stabilization in spinal cord injury. It is important to stress that microtubules need to be dynamic to support axon extension. Indeed, low concentrations of taxol do not block microtubule dynamics, but suppress the rate and extent of microtubule shrinkage, yet still allow polymerization (Derry et al., 1995). In contrast, high concentrations of taxol suppress microtubule dynamics, and thereby axon elongation (Schwamborn and Puschel, 2004), and cause toxicity for cells, which would imbalance the recovering tissue. Hence, a proper range of drug and delivering methods should be investigated for assessing the effect of taxol on axonal regeneration in vivo.
Putative signaling mechanisms underlying microtubule disorganization
Peripheral and central injuries trigger axonal responses that are dependent on distinct extrinsic factors present in the CNS and PNS. Microtubules are regulated by external signals during axon elongation and guidance (Suter et al., 1998; Buck and Zheng, 2002; Dent and Gertler, 2003). In fact, the known inhibitory cues present in the CNS myelin and the glial scar, including myelin-associated glycoprotein (MAG), Nogo, OMgP (oligodendrocyte–myelin glycoprotein), and versican, converge downstream of their receptors onto signaling pathways of the Rho GTPases (Dubreuil et al., 2003; Fournier et al., 2003; Schweigreiter et al., 2004; Sivasankaran et al., 2004), which, in addition to the actin cytoskeleton, also affect microtubule stability and dynamics (Etienne-Manneville, 2004; Mimura et al., 2006). This suggests that extracellular inhibitory molecules could also induce changes in the dynamics and morphology of the axonal tip by affecting microtubule organization. Indeed, a recent report from Mimura et al. (2006) showed that MAG inhibits microtubule assembly and axon outgrowth in postnatal cerebellar granule neurons via Rho kinase activation. Interestingly, they identified CRMP-2 (collapsin response mediator protein-2), a microtubule-associated protein, as a target for microtubule disassembly both in vitro and in vivo after spinal cord injury. In addition to externally triggered responses induced by inhibitory factors, there are intrinsic determinants that confer growth competence. Lesioning the peripheral axonal branches of DRG neurons causes expression of a distinct set of genes, termed regeneration-associated genes, which are not upregulated after central lesioning (Ylera and Bradke, 2006). Interestingly, those sets of genes contain direct and indirect regulators of microtubule polymerization and stabilization, including tubulin isoforms, MAPs (microtubule-associated proteins), and arginase I (Miller et al., 1989; Moskowitz and Oblinger, 1995; Cai et al., 2002; Bosse et al., 2006), which may enhance the rapid regrowth of lesioned PNS axons. Notably, it has been demonstrated that interfering with the expression of microtubule binding proteins inhibits neurite outgrowth (Caceres et al., 1992). Together, our work establishes microtubules aside from the actin cytoskeleton as a major intracellular regulator of growth restraint after CNS lesioning.
Injured axon terminals' retraction bulbs in vivo are hallmarks of regeneration failure in the injured spinal cord as well as most of the other CNS neurodegenerative disorders, including traumatic brain injury, Alzheimer's disease, Parkinson's disease, and multiple sclerosis (Coleman, 2005). Hence, identification of the differences inherent to formation of retraction bulbs and advancing growth cones would provide essential insights to better address the causes of regeneration failure in the CNS. We believe that our results decipher a key intracellular mechanism that determines the formation and differential growth properties of growth cones and retraction bulbs. The future challenge will be to manipulate the microtubules of retraction bulbs in vivo to induce regrowth of injured CNS axons. We expect that stabilization of microtubules, alone or combined with other therapeutic interventions, will be a potential approach to enhance regeneration of injured CNS axons.
This work was supported by the Max Planck Society and additional grants from the Deutsche Forschungsgemeinschaft (SFB 391). F.B. is a recipient of a Career Development Award from the Human Frontier Science Program. A.E. was supported by the Marie Curie Association. F.B. and A.E. are members of the European Union-Research Training Network “Nervous System Repair” (RTN MRTN-CT-2003-504636). J.E. is a student of the Gulbenkian PhD Program in Biomedicine and is sponsored by the Portuguese Foundation for Science and Technology. We thank D. Büringer and M. Braun for assistance in using electron microscopy. We are indebted to Dr. F. Barr for sharing reagents and to Dr. J. R. Sanes for the gift of GFP-M and YFP-H transgenic mice. We are grateful to Drs. B. Garvalov, H. Thoenen, A. Borst, R. Klein, C. Brakebusch, G. Tavosanis, M. Hübener, T. Keck, F. Barr, B. Ylera, S. Tahirovic, and Mr. H. Witte for their critical reading of this manuscript.
- Correspondence should be addressed to Frank Bradke, Max Planck Institute of Neurobiology, Axonal Growth and Regeneration, Am Klopferspitz 18, 82152 Martinsried, Germany.