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. 2009 May 20;29(20):6593–6598. doi: 10.1523/JNEUROSCI.1115-09.2009

PHR Regulates Growth Cone Pausing at Intermediate Targets through Microtubule Disassembly

Michael Hendricks 1, Suresh Jesuthasan 1,2,✉
PMCID: PMC6665902  PMID: 19458229

Abstract

Axonal growth cones use intermediate targets to navigate in the developing nervous system. Encountering these sites is correlated with growth cone pausing. PHR (Phr1, Esrom, Highwire, RPM-1) is a large neuronal ubiquitin ligase that interacts with multiple signaling pathways. Mouse and zebrafish phr mutants have highly penetrant axon pathfinding defects at intermediate targets. Mouse phr mutants contain excessive microtubules in the growth cone, which has been attributed to upregulation of DLK/p38 signaling. Here, we ask whether this pathway and microtubule misregulation are indeed linked to guidance errors in the vertebrate brain, using the zebrafish. By live imaging, we show that loops form when microtubules retract without depolymerizing. JNK, but not p38, phosphorylation is increased in mutant growth cones. However microtubule looping cannot be suppressed by inhibiting JNK. The phr microtubule defect can be phenocopied by taxol, while microtubule destabilization in vitro using nocodazole prevents loop formation. Acute disruption in vivo with nocodazole suppresses the intermediate target guidance defect. Given that microtubule looping is associated with growth cone pausing, we propose that microtubule disassembly, mediated by PHR, is essential for exiting the paused state at intermediate targets.

Introduction

Growing axons follow long paths that are broken up into smaller segments by intermediate targets. Typically, growth cones slow down or pause at these sites, while undergoing changes in morphology and signaling properties. Recent studies have described defects in axon pathfinding at intermediate targets, caused by mutations affecting microtubules and their associated proteins (Rothenberg et al., 2003; Lee et al., 2004, 2007), indicating that many guidance cues acting at intermediate targets regulate the microtubule cytoskeleton.

The PHR family of proteins includes human PAM, mouse Phr1, zebrafish Esrom, Drosophila Highwire, and Caenorhabditis elegans RPM-1 (Guo et al., 1998; Schaefer et al., 2000; Wan et al., 2000; Zhen et al., 2000; Burgess et al., 2004; D'Souza et al., 2005); for clarity we will refer to these as PHR. In vertebrates, phr mutants show highly specific axon pathfinding errors, which can be described as a failure to negotiate certain intermediate targets (Bloom et al., 2007; Lewcock et al., 2007; Hendricks et al., 2008). PHR is a huge, multidomain protein, and in invertebrates has been shown to regulate synapse formation via dual leucine zipper-bearing kinase (DLK), a member of the mixed lineage kinase (MLK) family of MAP3Ks, through its ubiquitin ligase activity (Nakata et al., 2005; Collins et al., 2006). In C. elegans, PHR is required for DLK-mediated activation of p38 MAP kinase (Nakata et al., 2005), whereas in Drosophila DLK acts via c-Jun N-terminal kinase (JNK) (Collins et al., 2006).

Cultured spinal neurons from phr mutant mice display growth cones with abnormal morphologies and disorganized microtubules (Lewcock et al., 2007). Since DLK was found away from the axon tip in phr mutants, and inhibition of p38 or treatment with taxol reduced microtubule disorganization, it was proposed that PHR acts outside of growth cones to stabilize microtubules by mediating destruction of DLK. However, loss of DLK fails to suppress axon pathfinding defects in the phr mutant mouse, and no increase in DLK was detected in phr mutants (Bloom et al., 2007). Furthermore, misregulation of p38 has not been demonstrated in PHR mutants. It is thus unclear whether and how the microtubule phenotype is linked to the pathfinding defect, or indeed, if microtubule regulation by PHR is mediated by DLK/p38 signaling.

The zebrafish phr mutant has a striking pathfinding defect in the dorsal forebrain (Hendricks et al., 2008). Axons that are destined to form the habenular commissure navigate normally to an intermediate target adjacent to the roof plate, but stall indefinitely and fail to cross the midline. Here, we use the zebrafish to test whether disorganized microtubules underlie the guidance defect at intermediate targets in phr mutants, and whether this occurs via overactivation of p38 or JNK.

Materials and Methods

Forebrain cultures.

Primary forebrain culture was performed as described previously (Hendricks and Jesuthasan, 2007). Microtubule immunofluorescence was based on Mitchison Lab protocols (http://mitchison.med.harvard.edu/protocols/gen1.html). Cultures were fixed for 10 min in microtubule fix [0.5% glutaraldehyde, 0.1% Triton X-100, and BRB80 (80 mm PIPES, pH 6.8, 1 mm MgCl2, and 1 mm EGTA)] and quenched with 0.1% sodium borohydrate for 7 min. For phospho-p38, fixation was done in 4% PFA, 20% sucrose, and PBS for 20 min at 4°C. For phospho-JNK and phospho-cJUN label, a 10 min fixation was performed in microtubule fix supplemented with 3.7% formaldehyde. Details on antibodies used are provided in the supplemental material (available at www.jneurosci.org).

Electroporation.

pNBT:EB3-EGFP, which contains the neuronal β-tubulin promoter driving EGFP-EB3 fusion (gift from Vincent Cunliffe, University of Sheffield, Sheffield, UK), was injected into the forebrain. Electrodes (PBSA0575, FHC) were used to apply several 30 V, 1 ms pulses from a Grass (Warwick) Telefactor SD9 voltage stimulator (Hendricks and Jesuthasan, 2007).

Pharmacology.

Stock solutions of SP600125 (Tocris) and SB203580 (Merck), were prepared at a concentration of 100 and 50 mm respectively, in cell culture grade DMSO (Sigma). Both compounds have previously been used on zebrafish (Fujii et al., 2000; Zhu et al., 2008).

Brains were injected with glass capillary needles made using a Sutter needle puller (Novato). Embryos at 2 dpf were mounted in agarose and a Narishige air pressure injector was used to deliver several picoliters of 0.33 μm nocodazole (in Ringers/10% DMSO) into the forebrain. The following day, embryos were fixed in 2% trichloroacetic acid/1% Triton X-100/PBS, washed, and stained with a monoclonal anti-acetylated tubulin antibody (1:1000, Sigma).

Imaging.

Cultures were imaged live on a Zeiss Axiovert microscope with a 100× phase objective (NA 1.4, Zeiss). Time-lapse images were acquired every 3 s with a Photometrics Cascade CCD camera using MetaMorph software. Fixed cultures were imaged on an inverted Zeiss LSM510 Meta confocal with a 63× oil-immersion lens. Whole mounts were imaged on either an upright Zeiss LSM510 or Exciter confocal using a 40× water-dipping objective.

Details on image analysis are provided in the supplemental material (available at www.jneurosci.org).

Results

Microtubules in phr mutant axons have an overgrowth phenotype

When forebrain neurons from zebrafish phr mutants were placed in culture, heterogeneous microtubule defects were observed in many axons (Fig. 1 A,B). These included single or clustered loops along the axon shaft, as well as complex snarls within growth cones.

Figure 1.

Figure 1.

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Microtubules in phr mutant axons. A, B, Forebrain explants from wild-type and phr embryos, α-tubulin immunofluorescence. Areas of insets (A′, B′) are shown with dashed lines. Microtubules in mutants exhibit small single or double loops (arrows) as well as large tangles (arrowheads). Kymograph analysis shows more rapid translocation of EB3-EGFP-positive microtubule plus-ends in mutants (D) compared with wild type (C) (Mann–Whitney U test, p < 0.01), as indicated by the slope of the puncta. E, Plot of all measured puncta showing quartile divisions of plus-end growth rate (slope), heavy bars show mean values. F, Frames from a time-lapse recording of wild-type axons with a moderate level of EB3 label. Microtubules grow and shrink (arrowheads indicate tips), without coil formation. This behavior is reflected in the kymograph (H) made from another wild-type axon. G, Mutant axons. The microtubule coils proximally (arrowhead) while the tip remains relatively stationary. I, Kymograph of a different mutant axon, again showing reduced shrinkage. A coil formed at the point indicated by the upper arrow. The microtubule retracted at the point indicated by the lower arrow. See supplemental material (available at www.jneurosci.org). In kymographs, d indicates distance and t indicates time. Scale bar, 20 μm.

To examine the mechanistic underpinnings of these defects, we performed time-lapse recordings of axons expressing the plus-end binding protein EB3 fused to EGFP. Typical “comets” corresponding to growing microtubules were observed in wild-type and mutant axons. Kymographic analysis of time-lapse videos revealed that plus-end translocation was significantly more rapid in mutants (Fig. 1 C–E). Average time for growth, as measured by persistence of EB3 signal, was the same in mutant and wild type: 8.8 ± 0.5 and 8.4 ± 0.6 respectively (number of frames ± SE, n = 42). This implies that longer polymers were assembled in mutants.

Neurons with a moderate level of EB3-EGFP, such that the entire microtubule was dimly labeled while the growing tip was more strongly fluorescent, allowed us to observe microtubule loops forming. In wild-type neurons (n = 5), microtubules grew slowly and shrank rapidly (Fig. 1 F,H; supplemental Movie 1, available at www.jneurosci.org as supplemental material), characteristic features of dynamic instability (Mitchison and Kirschner, 1984). In mutants (n = 6) with a similar or lower level of expression, growth was sometimes followed by little or no retraction, and this was accompanied by buckling and looping of the microtubule filament proximal to the plus-end (Fig. 1 G,I; supplemental Movie 2, available at www.jneurosci.org as supplemental material). Microtubules that formed subsequently polymerized along these loops.

JNK signaling is overactivated in phr mutant growth cones

Increased signaling via p38 has been proposed to cause the microtubules defects in mouse phr mutants. To test whether this pathway is overactivated in zebrafish mutants, we labeled cultured forebrain neurons with an antibody against active p38. No increase in label was observed in growth cones of mutants, compared with wild-type siblings (Fig. 2 A,B,D). There was no correlation between the level of phospho-p38 and the presence of microtubule defects (Fig. 2 C,D). Treatment of cultured axons with the p38 inhibitor SB203580 did not prevent the formation of microtubule loops (Fig. 2 E).

Figure 2.

Figure 2.

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Phosphorylation of p38 and JNK in phr mutants. A, B, Cultured wild-type and mutant forebrain axons labeled with anti-α-tubulin, DTAF (total protein), and anti-phospho-p38. C, Higher magnification of a mutant axon, showing an example of regions of analysis used for measuring the phospho-p38:DTAF ratio in normal (dashed line) versus defective (dotted line) microtubule areas. D, The proportion of phospho-p38:DTAF does not very between mutant and wild-type axons, and is the same in looped/disorganized regions as in adjacent areas of the axon. Bars indicate SEM. E, Mutant neurons after treatment with 50 μm SB203580. Microtubules, labeled with anti-α-tubulin, form loops in the axon. F, G, Wild-type and mutant forebrain neurons labeled with anti-α-tubulin (magenta) and anti-phospho-JNK (yellow). Arrows indicate growth axon tips with visible increase in phospho-JNK immunofluorescence. H, Ratio of phospho-JNK to DTAF in wild-type and mutant growth cones The plot shows quartile divisions of values and heavy bars indicate mean values. I, phr mutant axons, labeled for α-tubulin, following treatment with 20 μm SP600125. Loops are indicated by arrowheads. J, K, Assessment of SP600125 efficacy by analysis of phosphorylation of cJUN, a target of JNK. Levels of phospho-cJUN are high in control explants (0.1% DMSO; J), and lower in explants treated with 20 μm SP600125 (K). Images are pseudocolored, with white representing the highest intensity, as indicated by the wedge.

We next examined JNK, another MAP kinase that may be regulated by PHR. In contrast to p38, levels of phospho-JNK were elevated in mutant growth cones (Fig. 2 F–H). However, treatment of cultures with 2, 4, 10, or 20 μm SP600125 did not prevent loop formation (Fig. 2 I). These concentrations caused a reduction in phosphorylated c-JUN (Fig. 2 J,K), confirming that the drug was inhibiting JNK. Hence, upregulation of JNK in the phr mutant does not appear to be a critical factor in the formation of microtubule loops.

phr defects are phenocopied by taxol and suppressed by nocodazole

Live imaging of microtubules suggested that increased polymerization and/or stability might contribute to the microtubule defects in phr mutants. Consistent with this, treatment of wild-type forebrain explants with taxol, a compound that can increase the rate of tubulin polymerization in cells (De Brabander et al., 1981) and stabilize microtubules, resulted in the formation of numerous loops and knotted microtubule structures within axons and growth cones (Fig. 3 C,D,G). Microtubules were then examined for detyrosination (Fig. 3 H–K), a posttranslational modification that occurs gradually and is thus normally found in stable microtubules, e.g., primarily in the axon shaft. Twenty-one of 189 mutant axon tips contained this marker, in contrast to only two of 120 wild-type axons tips, indicating that a small proportion of mutant growth cones contain stabilized microtubules.

Figure 3.

Figure 3.

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Phenocopy and suppression of microtubule defects. A, B, Control cultures treated with DMSO (n wt = 276, nphr = 150). C, D, Overnight treatment with 20 nm taxol phenocopies the microtubule looping seen in phr (n wt = 156, nphr = 106). E, F, Nocodazole treatment leads to a reduction in microtubule defects (n wt = 71, nphr = 78). G, Proportion of axons showing one or more microtubule defects (single or multiple loops) (*χ2 test, p < 0.001). H–K, Double label for total (H, J) and detyrosinated tubulin. Regions containing detyrosinated tubulin are indicated in white in panels I and K. Arrowheads indicate axon tips containing low levels of detyrosinated tubulin, whereas arrows indicate axon tips with high levels of detyrosinated tubulin.

When treated with 20 nm nocodazole, which inhibits assembly of tubulin polymers and promotes microtubule depolymerization (Samson et al., 1979), phr cultures showed significant reduction in the proportion of axons displaying microtubule abnormalities (Fig. 3 E–G); this treatment had no effect on wild-type axons. Higher concentrations (50 nm) prevented axon outgrowth, while lower concentrations (10 nm) had no effect (data not shown).

Nocodazole suppresses habenular commissure defects

To test whether abnormal microtubule stability underlies the defect in habenular commissure formation, we treated mutants with acute doses of nocodazole at 48–52 hpf, when many axons have reached the intermediate target and stalled. Nocodazole was injected directly into the forebrain, and the following day embryos were assayed for commissure formation (Fig. 4). Control mutant siblings showed midline crossing in ∼10% of cases, consistent with the previously reported penetrance of the allele used here, phrtg06 (Hendricks et al., 2008). Approximately one-third of mutants injected with nocodazole (15/43) showed phenotypic suppression. Results were significantly different in two independent experiments (n = 15 and n = 28, χ2 test, p < 0.001). This suggests that PHR's role in regulating the microtubule cytoskeleton is essential for guidance at the midline.

Figure 4.

Figure 4.

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Nocodazole rescues the habenular commissure defect in phr mutants. A, Habenular commissure (arrow) of a 3 dpf wild-type embryo labeled for acetylated tubulin. B, The commissure is absent in phr mutants. C, Rescue of habenular commissure formation in an embryo injected with nocodazole at 2 dpf. Scale bar, 20 μm.

Discussion

Microtubules and intermediate targets

Growth cones often pause at intermediate targets, becoming larger and less polarized. Enlarged, paused growth cones in vitro as well as those at intermediate targets in vivo are characterized by an increase in microtubules and microtubule loops (Tsui et al., 1984; Sabry et al., 1991; Tanaka and Kirschner, 1991; Suh et al., 2004). This increase can occur when plus-ends are overstabilized, for example by overexpression of the tip-binding protein CLASP, or when the destabilizing protein SCG10 is inhibited, and this slows growth cone advance (Lee et al., 2004; Suh et al., 2004). Microtubule looping thus appears to be a cause, rather than an effect, of growth cone pausing.

Within growth cones, microtubules are variably coupled to retrograde actin flow (Forscher and Smith, 1988; Schaefer et al., 2002). The force produced by actin movements can cause microtubules to fold, buckle, or break, and this compression is a mechanism for loop formation (Tanaka and Kirschner, 1991). Recent work suggests the extent of microtubule retrotranslocation by actin can regulate growth cone advance. As coupling or local retrograde flow is decreased, microtubules can extend peripherally and stabilize nascent growth cone processes (Lee and Suter, 2008; Schaefer et al., 2008). If this model is correct, then tight coupling of microtubules to retrograde actin flow may be a mechanism to induce looping, and therefore pausing, at sites such as intermediate targets.

phr mutants and the tubulin cytoskeleton

In zebrafish phr mutants, forebrain axons exhibit extensive microtubule looping, similar to those observed in mouse spinal neurons in vitro (Lewcock et al., 2007). The formation of loops and growth cones with knotty microtubule arrays has been described in neurons treated with taxol (Letourneau and Ressler, 1984; Williamson et al., 1996; Rajnicek et al., 2006). Chronic treatment with a low dose of taxol indeed phenocopies these aspects of the phr phenotype in our forebrain cultures. To the contrary, Lewcock et al. (2007) observed a reduction in growth cone microtubules and their disorganization using a similar taxol treatment. This difference may have occurred because the effects of taxol vary according to concentration, and the effective dose differs according to culture medium, culturing temperature, species, and cell type (see for example Yvon et al., 1999).

Despite the prevalence of microtubule defects in cultured phr neurons, the in vivo axon guidance phenotypes are highly specific and most axonal projections are normal. For this reason, it is unlikely that phr growth cones have general defects in chemotaxis or motility. We suggest that when growth cones encounter sites where microtubule regulation is critical, pathfinding fails. At the roof plate boundary in the zebrafish diencephalon, a Wnt-expressing intermediate target for commissural axons, growth cones normally pause and enlarge before crossing. Wnt can regulate axonal microtubules and induce growth cone enlargement via multiple pathways (Gordon-Weeks, 2004; Ciani and Salinas, 2007; Purro et al., 2008; Salinas and Zou, 2008). We suggest that the additive effects of Wnt signaling at the intermediate target and increased microtubule polymerization and stability in phr mutants cause excessive microtubule looping, preventing growth cones from exiting the paused state.

How does PHR regulate microtubules?

In Drosophila and C. elegans, PHR inhibits a conserved MAP kinase cascade consisting of DLK, MKK4, and p38 or JNK (Nakata et al., 2005; Collins et al., 2006). In the zebrafish, levels of phospho-JNK are visibly increased in mutants. However, JNK does not appear to be involved in the formation of loops, as inhibition with SP600125 does not prevent loop formation. This is in agreement with Bloom et al. (2007), who find that loss of DLK, which can activate JNK, cannot suppress the guidance phenotype in mouse phr mutants. Lewcock et al. (2007) have suggested that PHR regulates microtubules via p38, downstream of DLK. We find no evidence of increased p38 signaling in the zebrafish phr mutant. This may reflect species or tissue differences; alternatively, the reduction in microtubules caused by p38 inhibition in mouse phr growth cones may have been caused by inhibition of a parallel pathway. Consistent with this, treatment of wild-type mouse growth cones with SB203580 caused a reduction in microtubules (Lewcock et al., 2007), and treatment of the mutant only resulted in a partial reduction.

PHR may function within a complex that directly interacts with the cytoskeleton. Intriguingly, the N terminus of PHR shows high homology to the yeast ATS1 (α-tubulin suppressor 1) protein, which reduces microtubule levels through an unknown mechanism (Kirkpatrick and Solomon, 1994). Lewcock et al. (2007) demonstrated that PHR remains associated with the microtubule cytoskeleton after detergent extraction, and recent work by Pierre et al. (2008) shows association with actin. While the former study suggested that PHR is excluded from the growth cone and functions in the axon, the latter shows growth cone localization in mouse spinal neurons. We also observed PHR in the growth cones of both retinal ganglion cells and forebrain neurons in zebrafish, by immunofluorescence (D'Souza et al., 2005; Hendricks et al., 2008).

We hypothesize that PHR acts in the growth cone to regulate microtubule polymerization and stability. This is in contrast to the model of Lewcock et al. (2007), which proposes that PHR is excluded from the growth cone, and in its absence DLK is activated away from the axon tip. We suggest that in the absence of PHR, microtubule dynamics are biased toward growth and may be excessively coupled to actin flow, causing inappropriate looping and a failure to exit a paused state. As predicted by this model, these microtubule defects are phenocopied by taxol and suppressed by nocodazole. Most strikingly, nocodazole rescues growth cone stalling at an intermediate target, constituting the first in vivo suppression of phr axon guidance defects. In summary, our results demonstrate that microtubule disorganization causes the phr guidance phenotype, provide further evidence that microtubule polymerization is causal to growth cone pausing, and suggest that PHR-mediated regulation of microtubule stability is critical to pathfinding at intermediate targets.

Footnotes

This work was funded by the Singapore Millennium Foundation and the Biomedical Research Council. We thank Erik Dent and Ajay Mathuru for comments on the manuscript.

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