Cover ArticleResearch Articles, Development/Plasticity/Repair

The MAP3Ks DLK and LZK Direct Diverse Responses to Axon Damage in Zebrafish Peripheral Neurons

Kadidia Pemba Adula, Matthew Shorey, Vasudha Chauhan, Khaled Nassman, Shu-Fan Chen, Melissa M. Rolls and Alvaro Sagasti
Journal of Neuroscience 10 August 2022, 42 (32) 6195-6210; DOI: https://doi.org/10.1523/JNEUROSCI.1395-21.2022

Abstract

Mitogen-activated protein kinase kinase kinases (MAP3Ks) dual leucine kinase (DLK) and leucine zipper kinase (LZK) are essential mediators of axon damage responses, but their responses are varied, complex, and incompletely understood. To characterize their functions in axon injury, we generated zebrafish mutants of each gene, labeled motor neurons (MNs) and touch-sensing neurons in live zebrafish, precisely cut their axons with a laser, and assessed the ability of mutant axons to regenerate in larvae, before sex is apparent in zebrafish. DLK and LZK were required redundantly and cell autonomously for axon regeneration in MNs but not in larval Rohon–Beard (RB) or adult dorsal root ganglion (DRG) sensory neurons. Surprisingly, in dlk lzk double mutants, the spared branches of wounded RB axons grew excessively, suggesting that these kinases inhibit regenerative sprouting in damaged axons. Uninjured trigeminal sensory axons also grew excessively in mutants when neighboring neurons were ablated, indicating that these MAP3Ks are general inhibitors of sensory axon growth. These results demonstrate that zebrafish DLK and LZK promote diverse injury responses, depending on the neuronal cell identity and type of axonal injury.

SIGNIFICANCE STATEMENT The MAP3Ks DLK and LZK are damage sensors that promote diverse outcomes to neuronal injury, including axon regeneration. Understanding their context-specific functions is a prerequisite to considering these kinases as therapeutic targets. To investigate DLK and LZK cell-type-specific functions, we created zebrafish mutants in each gene. Using mosaic cell labeling and precise laser injury we found that both proteins were required for axon regeneration in motor neurons but, unexpectedly, were not required for axon regeneration in Rohon–Beard or DRG sensory neurons and negatively regulated sprouting in the spared axons of touch-sensing neurons. These findings emphasize that animals have evolved distinct mechanisms to regulate injury site regeneration and collateral sprouting, and identify differential roles for DLK and LZK in these processes.

Introduction

Axon damage caused by stroke, trauma, or disease disrupts the circuits required for sensation, movement, and cognition. Unlike other tissues that rely on stem cells to recover from damage, most neurons cannot be replaced, so damaged cells must themselves be repaired to restore function. Successful axon regeneration requires damage sensing, the transmission of injury signals to the nucleus, activation of proregenerative genes, axon guidance, and circuit reintegration (Curcio and Bradke, 2018). Our understanding of the factors regulating each of these steps is incomplete.

Mitogen-activated protein kinase kinase kinases (MAP3Ks) regulate many cellular processes, including development, differentiation, and stress responses (Craig et al., 2008). Among MAP3Ks, dual leucine kinase (DLK/MAP3K12), which belongs to the mixed lineage kinase (MLK) MAP3K subfamily (Gallo and Johnson, 2002), has been implicated in neuronal development (Nakata et al., 2005; Hirai et al., 2006, 2011) and axon injury responses (Hammarlund et al., 2009; Miller et al., 2009; Xiong et al., 2010; Welsbie et al., 2017, 2019; Saikia et al., 2022) in a variety of organisms. DLK is activated by axon injury and in turn activates downstream P38 or JNK signaling cascades to induce transcription of injury response genes (Tedeschi and Bradke, 2013; Jin and Zheng, 2019). In addition to DLK itself, vertebrates have another DLK-related gene that contributes to injury responses, leucine zipper kinase (LZK/MAP3K13), which has a domain analogous to the calcium-sensing domain in the worm DLK protein (Yan and Jin, 2012).

In invertebrates, activation of DLK is a major signal initiating responses to axon injury and stress. Without DLK, regenerative axon growth is eliminated in motor and sensory neurons in both Caenorhabditis elegans (Hammarlund et al., 2009; Yan et al., 2009) and Drosophila (Xiong et al., 2010; Stone et al., 2014). In mammals, the outcomes of DLK and LZK activation in response to axon damage are varied and context dependent. DLK or LZK can promote neurite branching (Chen et al., 2016), axon elongation (Shin et al., 2012), axon regeneration in PTEN (phosphatase and tensin) mutants (Saikia et al., 2022), injury-induced axon sprouting (Saikia et al., 2022), inhibition of axon regeneration (Dickson et al., 2010), axon degeneration of the distal stump (Miller et al., 2009; Summers et al., 2018), cell death (Ghosh et al., 2011; Watkins et al., 2013; Yin et al., 2017; Welsbie et al., 2019; Li et al., 2021), or microglial and astrocyte responses to injury (Chen et al., 2018; Wlaschin et al., 2018). In mice with a DLK gene trap, downstream responses were reduced after sciatic nerve injury, and explant cultures of dorsal root ganglion (DRG) grew shorter axons than controls (Itoh et al., 2009). Selectively eliminating DLK in neurons strongly reduced motor axon target reinnervation after sciatic nerve crush (Shin et al., 2012). Peripheral axons of DLK mutant DRG neurons, on the other hand, initiated regeneration normally but by 3 d postinjury had regenerated less than controls (Shin et al., 2012).

Studying genetic regulators of axon regeneration is complicated by the complexity of in vivo experimental injuries. In nerve crush or severing models, nerve bundles contain axons of many different types of neurons, thus convoluting cell-type-specific responses to injury. Moreover, axon branching and fasciculation within nerves can obscure the source of growth after incomplete injuries; without single cell resolution, it is difficult to distinguish axon growth at the site of injury from growth of spared branches in a damaged neuron or from collateral sprouting of undamaged neurons. Here, we refer to growth from unsevered branches of injured cells as “regenerative sprouting” and compensatory growth from neurons that have not been directly damaged as “collateral sprouting,” similar to how others have defined these terms (Steward et al., 2003; Tuszynski and Steward, 2012). Distinguishing between each of these kinds of axon growth is important because each has distinct implications for the functional restoration of neuronal circuits.

To better understand the roles of DLK and LZK in axon damage responses, we created zebrafish mutants in the genes encoding both proteins. Single-cell labeling allowed us to compare regeneration not only in different cell types but also in central axons, peripheral axons, and specific branches of damaged axons. Our findings indicate that DLK and LZK are required redundantly for motor axon regeneration but not for axon regeneration in Rohon–Beard (RB) or DRG neurons. Surprisingly, DLK inhibited sprouting of larval peripheral sensory axons, emphasizing the context-dependent multifunctionality of these axon damage sensors.

Materials and Methods

Zebrafish

Zebrafish (Danio rerio) were raised on a 14/10 h light/dark cycle and a water temperature of 28.5°C. Embryos at early larval stages were used before sex is apparent. Thus, both presumptive male and female embryos were used for all experiments. Embryos were incubated at 28.5°C in E3 buffer (0.3 × g/l Instant Ocean salt, 0.1% methylene blue). For imaging purposes, pigment formation was blocked by treating embryos with phenylthiourea (PTU; 1 × PTU, 0.2 mm) at 22–24 h post-fertilization (hpf). Embryos were then manually dechorionated using forceps. All mutant and transgenic lines were created using AB wild-type fish (ZFIN: ZDB-GENO-960809–7). Experimental procedures were approved by the Chancellor's Animal Research Care Committee at University of California, Los Angeles and the Pennsylvania State Institutional Animal Care and Use Committee.

CRISPR/Cas9 mutagenesis

Guide RNAs were engineered using the short oligo method to generate gRNAs, as previously described (Talbot and Amacher, 2014). To mutagenize dlk and lzk, we created a DNA template for making gRNAs, containing the T7 RNA polymerase promoter, the gene targeting sequence, and the gRNA scaffold sequence. This template was amplified by PCR, and its product was used to synthesize gRNAs using a T7 RNA polymerase kit. The gRNAs were then purified. Fish were injected with a mix containing 1 ul of Cas9 mRNA (1500 ng/ul), 1.5 ul of gRNA1 (100 ng/ul), and 1.5 ul of gRNA2 (100 ng/ul). Embryos at the one-cell stage were injected with 5 nl of the mix. At 48 hpf, PCR and restriction digests were used to test guide efficiency. To identify fish carrying mutations in their germline, injected embryos were raised to adulthood, crossed to wild-type fish, and their progeny were screened by PCR. At 48 hpf, mutant PCR products flanked by the common M13 primers were sent for sequencing.

PCR genotyping

PCR was usually conducted with Taq polymerase for 40 amplification cycles. The denaturation cycle was 94°C for 30 s. Annealing was for 30 s at 53°C for lzk wild-type and lzkla232 bands, 59°C for the dlk wild-type band, and 63°C for dlkla231 band. Elongation was at 72°C; this step was 20 s for lzk wild-type and lzkla232 bands, 1 min for the dlk wild-type band, and 15 s for the dlkla231 band.

Transgene cloning

HB9(3×)-E1B-DLK-T2A-GFP

E1B-DLK-T2A was constructed by individually inserting E1B and DLK into the MCS region of a PME: MCS-T2A vector.

Step 1, E1B template

5′-TCTAGAGGGTATATAATGGATCCCATCGCGTCTCAGCCTCA-3′

5′-GAATTCGTGTGGAGGAGCTCAAAGTGAGGCTGAGACGCGATG-3′ was created by PCR amplification of overlapping oligomers.

Step 2 was att site-MCS-T2A-att site

5′-GGGGACAAGTTTGTACAAAAAAGCAGGCTACCGTCAGATCCGCTAG-3′

5′-GGGGACCACTTTGTACAAGAAAGCTGGGTATGGGCCAGGATTCTC-3′. MCS-T2A flanked by att sites was PCR amplified and inserted into a PME vector using a Gateway BP reaction (Kwan et al., 2007).

In Step 3, EcorI-E1B-EcorI was inserted into the MCS region of PME: MCS-T2A vector using restriction digest and ligation resulting in aPME: MCS(E1B)-T2A vector

5′-TAAGCAGAATTCTCTAGAGGGTATATAATGGATCCCA-3′

5′-TGCTTAGAATTCGAATTCGTGTGGAGGAGCT-3′.

In Step 4, SaLI-DLK(no stop codon)-SacII was inserted into the PME: MCS(E1B)-T2A vector using restriction digest and ligation resulting in a PME: MCS(E1B-DLK)-T2A vector

5′-TAAGCAGTCGACATGGCTTGTGTCCATGAGCAG-3′

5′-TGCTTACCGCGGGTTTTGTGGACCCTGGCCC-3′. PME: MCS(E1B-DLK)-T2A, a P5E:HB9(3×), and P3E:GFP were incorporated into a Gateway destination vector via LR reaction resulting in HB9(3×)-E1B-DLK-T2A-GFP.

HB9(3×)-E1B-LZK-T2A-GFP

E1B-LZK-T2A was constructed using overlap PCR to assemble E1B-LZK-T2A framed by att sites as follows: Primer set 1, att site-E1B-part of lzk, use E1B sequence as a template

5′-GGGGACAAGTTTGTACAAAAAAGCAGGCTTCTCTAGAGGGTATATAATGGATCCC-3′

5′-TGGTGCTGTGCGTGTGCATGAATTCGTGTGGAGGAGC-3′. Primer set 2, part of E1B-lzk no stop codon-part of T2A, use lzk cDNA as template

5′-GCTCCTCCACACGAATTCATGCACACGCACAGCACCA-3′

5′-CCTCTGCCCTCTCCACTTCCCCAGGATGACGGAGCGCC-3′. Primer set 3, end of lzk no stop codon-T2A-att site, use T2A sequence as a template

5′-GGCGCTCCGTCATCCTGGGGAAGTGGAGAGGGCAGAGG-3′

5′-GGGGACCACTTTGTACAAGAAAGCTGGGTCTGGGCCAGGATTCTCCTCGA-3′.

All three fragments were amplified independently. Fragment 1 was added to fragment 2 by overlapping PCR. The resulting fragment was added to fragment 3. The complete sequence was then inserted into the Gateway PME donor plasmid via a BP reaction. The resulting PME: E1B-LZK-T2A, a P5E:HB9(3×), and P3E:GFP were incorporated into a Gateway destination vector via LR reaction resulting in HB9(3×)-E1B-LZK-T2A-GFP.

Crest3: DLK-T2A-GFP

DLK-T2A was constructed using overlap PCR to assemble DLK-T2A flanked by att sites for Gateway recombination as follows: Primer set 1, att site-dlk no stop-part of T2A, use dlk cDNA as template

5′-GGGGACAAGTTTGTACAAAAAAGCAGGCTTCATGGCTTGTGTCCATGAGCAG-3′

5′-CCTCTGCCCTCTCCACTTCCGTTTTGTGGACCCTGGCCC-3′. Primer set 2, end of dlk no stop-T2A-att site, use T2A sequence as template

5′-GGGCCAGGGTCCACAAAACGGAAGTGGAGAGGGCAGAGG-3′

5′-GGGGACCACTTTGTACAAGAAAGCTGGGTCTGGGCCAGGATTCTCCTCGA-3′.

Both fragments were amplified independently. Fragment 1 was added to fragment 2 by overlapping PCR. The resulting PME: DLK (no stop codon)-T2A, a P5E: Crest3, and a P3E:GFP were incorporated into a Gateway destination vector via LR reaction resulting in Crest3: DLK-T2A-GFP.

Building the phylogenetic tree

Complete DLK and LZK protein sequences from several organisms were downloaded from the National Center for Biotechnology Information database. Accession numbers are shown in Table 1. The sequences were aligned using the MUSCLE alignment algorithm on the European Molecular Biology Laboratory–European Bioinformatics Institute website. Phylogenetic analyses were performed with RAxML (Randomized Axelerated Maximum Likelihood) software using a maximum likelihood method, the JTT substitution matrix, and empirical frequencies (Stamatakis, 2014). The RAxML software was accessed via the CIPRES (Cyberinfrastructure for Phylogenetic Research) Science Gateway (Miller et al., 2010). The Interactive Tree of Life website (Letunic and Bork, 2007) was used to visualize the evolutionary tree.

View this table:
Table 1.

Accession numbers

Larval body measurements

The body lengths of larvae of each genotype were measured at 48 days postfertilization (dpf), 72, and 5 dpf using a Zeiss Discovery V8 Stereomicroscope at a 2× magnification. Bodies were measured lengthwise from head to tail, and tail widths measured ventral to dorsal at the end of the spinal cord. Larvae from different clutches (n > 8) and parents of different ages (3–24 months old) were mixed for this analysis.

Mounting larvae for live imaging

Only one neuron per larva was tracked for imaging and axotomy. Following screening and identification of an isolated RB neuron or motor neuron (MN), each individual larva was grown in its own agarose-coated 35 mm dish or the well of a six-well plate.

Larva were anesthetized with 0.2 mg/ml MS-222 in embryo media (0.08%) before mounting. Each larva was embedded in 1% agarose and placed on a coverslip. On solidification of the agarose (∼15 min), a plastic ring was sealed onto the coverslip with vacuum grease. The resulting chamber was then filled with tricaine-containing embryo media and sealed with a glass slide using vacuum grease (O'Brien et al., 2009a).

Larval microscopy and axotomy

Live confocal images were collected on an LSM 800 using a 20× air objective (Plan-Apochromat, NA = 0.8). Images were acquired with Zen Blue software from Zeiss.

Laser axotomies were performed using a Zeiss LSM 880 confocal microscope equipped with a two-photon laser (Chameleon Ultra II, Coherent), as previously described (O'Brien et al., 2009a). Zen Black 2.1 SP3 software was used to visualize axons and perform axotomies. Neurons were visualized with a 561 nm or 488 nm laser excitation before switching to the Chameleon (813 nm) laser for axon severing.

To cut axons of MNs, we initially attempted axotomy with 6.5% laser power. If axons were not cut, the laser power was increased in 0.5% increments until the axon was severed. Successful axotomy was verified by observing a gap in axonal fluorescence immediately after laser illumination and detecting Wallerian degeneration of the distal segment several hours later. RB neurons were cut using a similar method, but we initially used 5% laser power to attempt axotomy because their more superficial location made them easier to cut.

Adult experiments

For cutting DRG nerves in juvenile fish, 4–5 week-old fish of the genotype P2rx3a:LexA,4xLexOP:EB3-GFP (data not shown), P2rx3a:LexA, 4xLexAop:mCherryla207, and Roya9/Roya9, miftaw2/miftaw2, containing either wt or homozygous dlkla231 and lzkla232 were anesthetized in a VWR polystyrene Petri dish, filled halfway with 0.16% tricaine in 0.6 × g/l Instant Ocean salt solution and immobilized by applying agarose to their midsection only, leaving both the respiratory apparatus of the head and tail free. Fish were then imaged on a Leica SP8 microscope equipped with an InSight X3 unit from Spectra-Physics. A 25× (NA = 1) water immersion objective with a working distance of 2.6 mm was used to image the posterior spinal cord of the fish, and a region of interest was chosen to restrict the cut site to the width of the nerve and positioned ∼100 μm from the posterior-most DRG, along the posterior projecting nerve. The tunable laser was set to 900 nm, and both the tunable and fixed wavelength 1045 nm lasers were set to 100% on the slowest speed setting and scanned for ∼1 s. The two-photon overview showing the nerve stumps after cut was performed on the aforementioned SP8, all other images for this experiment were obtained with a Zeiss LSM 800 Axio Observer Z1 with a 20× air objective (NA = 0.8).

In cutting scale nerves, all fish were between 8 and 11 months old. The single dlkla231 and lzkla232 mutants were transgenic for P2rx3a:LexA, 4xLexAop:mCherryla207 and did not possess mutant roy or mifta alleles. Wild-type and double mutant fish were in a roya9/roya9, miftaw2/miftaw2 background and doubly transgenic for both P2rx3a:LexA,4xLexAop:EB3-GFP and P2rx3a:LexA,4xLexAop:mCherryla207. All images shown were collected on a Zeiss LSM 800 Axio Observer Z1. For wild-type, a 25× a multi-immersion objective was used (NA = 0.8), and for the double mutants a 20× air objective (NA = 0.8) was used. Laser injury was performed using an Andor MicroPoint UV pulse laser.

Image analysis and statistics

Confocal images, saved as CZI files, were opened in ImageJ/Fiji and measured using the Simple Neurite Tracer feature in z-stack format. One neuron was imaged per embryo. Axons of MNs were traced from the cell body to their endings in muscles. RB peripheral arbors were traced starting at the first branch point in the skin. If RB peripheral axons bifurcated in the spinal cord, creating two separate peripheral arbors, both arbors were traced. Three experimenters contributed to tracing. To test for tracing reproducibility among experimenters, a subset of RB central axons (n = 76) and peripheral arbors (n = 10) were separately traced by two experimenters. In both cases, tracings by the two experimenters were highly similar (r = 0.93 for the central axons; r = 0.99 for the peripheral arbors).

For figures, maximum projections were created in ImageJ/Fiji, converted to grayscale, and inverted. To visualize entire RB neurons, images of different parts of each neuron were stitched together in Adobe Photoshop.

Dot plots or box-and-whisker plots overlaid with dot plots were produced in R software to visualize the data. In dot plots, the black bar indicates the mean. In box-and-whisker plots, the boxes indicate the interquartile range; the top of the box is the 75th percentile, the midline is the median, and the bottom of the box is the 25th percentile. Statistical analyses were performed in R. A generic quantile-quantile test was used to determine the normality of sample populations. Unless otherwise specified, data distributions were nonparametric. Therefore, a Kruskal–Wallis with a Bonferroni correction was used to assess differences between experimental groups. A Wilcoxon paired test was used to identify groups with significant differences (*p < 0.05, **p < 0.01, ***p < 0.005, and ****p < 0.001). Tables provide a comprehensive lists of genes, guide RNAs and primers (Table 2), plasmids and transgenic lines (Table 3), and other reagents and resources (Table 4).

View this table:
Table 2.

Gene IDs, Cas9 gRNAs, genotyping primers

View this table:
Table 3.

Plasmids used in injections, transgenic lines

View this table:
Table 4.

Reagents, resources

Results

dlkla231 and lzkla232 mutant zebrafish develop normal motor and sensory neurons

To study the functions of zebrafish DLK and LZK in axon regeneration, we identified the closest homologous genes to mammalian DLK and LZK in the zebrafish genome, which were located on opposite ends of chromosome 9, and generated a phylogenetic tree with the full amino acid sequences (Fig. 1A). Human, mouse, and zebrafish DLK proteins share 93% sequence similarity in their kinase domain and 95% similarity in their leucine zipper domains; human, mouse, and zebrafish LZK proteins share 97% sequence similarity in their kinase domain, 98% similarity in their leucine zipper domains, and an identical C-terminal hexapeptide involved in calcium regulation.

Figure 1.
Figure 1.

dlk And lzk zebrafish mutants. A, Phylogenetic tree of DLK and LZK orthologs. B, Genotyping of dlkla231 and lzkla232 CRISPR/Cas9 mutants. Left, gene structure of zebrafish dlk and lzk, with gRNA sequences. Blue indicates PAM (protospacer adjacent motif) sites. Right, DNA gel showing WT and mutant genotyping with primers indicated to the left (arrowheads). C, Forty-eight hpf zebrafish larvae of the indicated phenotypes. D, Overlaid box and dot plots comparing animal lengths from the tip of the head to the end of the tail each genotype. E, Overlaid box and dot plots comparing tail width in each genotype (see above, Materials and Methods; Table 5 for details of statistical analyses).

View this table:
Table 5.

p Values

We created mutations in each gene using the CRISPR/Cas9 system. Using two guide RNAs (gRNAs), we made a 3289 base pair (bp) genomic deletion in zebrafish dlk, which removed 519 coding base pairs, including most of the kinase domain. Using the same approach to target LZK, we created an allele with two separate deletions in exon 1 (a 28 bp deletion and a 30 bp deletion), placing the gene out of frame, upstream of the kinase domain (Fig. 1B). Unlike DLK mutant mice, which die perinatally (Hirai et al., 2006), dlkla231, lzkla232, and dlkla231 lzkla232 double mutant zebrafish survived to adulthood and appeared grossly normal, similar to mutants in invertebrate DLK homologs, although both mutants were slightly smaller on average than wild-type fish at three larval stages (48, 72, and 5 dpf), perhaps indicating a developmental delay (Fig. 1C–E).

Zebrafish dlk mRNA is expressed broadly in the nervous system at early developmental stages (Thisse et al., 2004), prompting us to test if dlk is required for the initial development of peripheral neurons. To label MNs, one-cell stage embryos were coinjected with MN driver (HB9:GAL4; Issa et al., 2011) and effector (UAS:GFP) transgenes, resulting in motor-neuron-specific expression of cytoplasmic GFP. As transient transgenesis results in mosaic inheritance, we screened for animals expressing GFP in isolated MNs. To minimize morphologic variability, we selected only those MNs that innervated ventral muscles (Fig. 2A,B), enriching for the caudal primary (CaP) MNs (Westerfield et al., 1986). At 5 dpf we imaged each MN and measured several morphologic parameters, including axon branch tip number, and total axon length. Despite the fact that mutant animals are slightly smaller than controls, there were no significant differences in neuronal morphology between dlkla231, lzkla232, or dlkla231 lzkla232 double mutants and wt animals (Fig. 2C,D). These observations indicate that dlk and lzk are not required for the morphologic development of MNs.

Figure 2.
Figure 2.

Motor neurons develop normally in dlkla231 and lzkla232 mutants. A, Diagram of 5 dpf larva, showing the approximate location of the image below of a single labeled MN in a live animal. The cell body and dendrites are in the spinal cord; the axon exits the spinal cord to innervate the ventral muscles of one segment. B, Labeled MNs in each of the indicated genotypes. C, Dot plot showing lengths of MNs in each of the indicated genotypes. Bar indicates the mean. There was no significant difference between groups (because distributions were normal, groups were compared by ANOVA). D, Dot plot showing branch tip numbers of MNs in each of the indicated genotypes. Bar indicates the mean. There was no significant difference between groups (see above, Materials and Methods; Table 5 for details of statistical analyses). Scale bars: A, 100 μm; B, 50 μm.

To image larval RB touch-sensing neurons (Palanca et al., 2013; Katz et al., 2021), we injected animals with a reporter driving expression of a red fluorescent protein in these cells (Isl1[SS]:Gal4;UAS:DsRed; Sagasti et al., 2005), and screened for animals expressing this reporter in isolated RB neurons, which allowed us to unambiguously visualize the morphology of the entire neuron and distinguish central from peripheral axons. To minimize variability, we selected only tail-innervating RB neurons for analysis because they are flat and relatively easy to trace (Fig. 3A,B). As with MNs, there were no significant morphologic differences between the peripheral arbors of dlkla231, lzkla232, or dlkla231 lzkla232 double mutant and wt fish at 48 hpf (Fig. 3C–E). At 72 hpf, dlkla231 and double mutant peripheral axon morphologies were also comparable to wild-type axons, but lzkla232 neurons were somewhat smaller than wild-type neurons on average, potentially reflecting their mild developmental delay. Together these results indicate that similar to homologs in invertebrate animals, DLK and LZK do not play major roles in initial neuronal development.

Figure 3.
Figure 3.

Rohon–Beard neurons develop normally in dlkla231 and lzkla232 mutants. A, Diagram of 48 hpf larva, showing the approximate location of the image below of a single labeled RB neuron in a live animal. The cell body and central and peripheral axons are labeled. The cell body and central axon are in the spinal cord; the peripheral axon exits the spinal cord to arborize in the developing epidermis. B, Tail-innervating peripheral RB axon arbors of the indicated genotypes at 48 and 72 hpf. C–E, Quantification of RB peripheral axon arbor length (C), 2D arbor area (D), and branch tip number (E) at 48 hpf (see above, Materials and Methods; Table 5 for details of statistical analyses). Scale bars: 100 μm.

dlk and lzk are redundantly required for motor axon regeneration

Few studies have directly tested the relationship between these two closely related, potentially redundant MAP3K proteins in axon damage responses. To assess if dlk or lzk are required for motor axon regeneration in larval zebrafish, we severed axons of isolated MNs in wt, single mutant, and double mutant fish and compared their ability to regenerate. Specifically, 5 dpf axons were severed 50 µm distal to the spinal cord exit point (Fig. 4A), using a laser mounted on a two-photon microscope (O'Brien et al., 2009a). To minimize potential contributions from extrinsic factors, only neurons in which laser axotomy caused no obvious damage or scars were used for regeneration experiments. Successful axotomy was verified by observing debris and a gap in fluorescence at the axotomy site and by confirming that disconnected distal branches later degenerated by Wallerian degeneration. Motor axons were assessed for regeneration 48 h postaxotomy (hpa; Fig. 4A), as in previous studies (Rosenberg et al., 2012). Wallerian degeneration appeared to occur normally in these mutants. Motor axon regeneration was modestly reduced in dlkla231 mutants, compared with wt motor axons, but strongly impaired in dlkla231 lzkla232 double mutants (Fig. 4B,C). These observations suggest that dlk and lzk are partially redundant (or genetically compensate for each other) for regeneration of larval motor axons.

Figure 4.
Figure 4.

Motor neuron regeneration is impaired in dlkla231 lzkla232 mutants. A, Top, Five days postfertilization motor axons immediately after axotomy in the indicated genotypes. Lightning bolt indicates axotomy site. Magenta highlights the separated distal stump that will degenerate. Far right, Neurons expressing rescue cDNAs; expression of rescue transgenes in cell bodies is shown below. Bottom, Same neurons 48 h postaxotomy. Blue highlights regenerated axons. B, C, Dot plots showing total regenerated length in each genotype (B) and the percentage of the original axon length regenerated (C). Bar indicates the mean. D, Dendrite overgrowth phenotype following failure of axon regeneration in dlkla231 lzkla232 mutants. E, Quantification of dendrite overgrowth (see above, Materials and Methods; Table 5 for details of statistical analyses). Scale bars: 50 μm.

Intriguingly, we observed that among dlkla231 lzkla232 double mutant MNs displaying severe regeneration deficits, ∼28% (8 of 29) grew extremely long neurites within the spinal cord (Fig. 4D,E), which are likely dendrites, a phenotype not seen in individual dlkla231 or lzkla232 mutants. This observation indicates that dlkla231 lzkla232 double mutant axons do not lack growth potential but are specifically impaired in axon regeneration.

To determine whether dlk or lzk are required cell autonomously for motor axon regeneration, we attempted to rescue their regeneration defects by expressing dlk and lzk cDNAs specifically in these neurons. As dlk and lzk were required redundantly for motor axon regeneration, we expressed each cDNA separately in dlkla231 lzkla232 double mutants. Strong overexpression of these cDNAs with the Gal4/UAS system was toxic to neurons, causing dysmorphic axons, spontaneous axon degeneration, and cell death (data not shown). We therefore expressed lower levels by creating bicistronic transgenes directly under an MN-specific promoter (HB9:DLK-T2A-GFP and HB9:LZK-T2A-GFP), which were coinjected with transgenes that strongly drive RFP expression throughout the cytoplasm (HB9:Gal4 and UAS:DsRed). Only cells that clearly expressed the rescue transgene (i.e., GFP+ cells), and had overtly normal axons, were used for these experiments (Fig. 4A). Expressing each cDNA improved axon regeneration to levels comparable to the corresponding single mutant (e.g., dlkla231 lzkla232 neurons with dlk cDNA were comparable to lzkla232 mutants; Fig. 4B,C). However, the difference in regeneration between rescued neurons and double mutants did not reach significance, likely because of variability in the assay. Nonetheless, these results are most consistent with DLK and LZK proteins acting cell autonomously to promote motor axon regeneration, similar to the well-documented role of DLK in axon regeneration in worms, flies, and mice.

dlk and lzk are not required for RB central or peripheral axon regeneration

RB neurons are bipolar or pseudo unipolar, with a receptive peripheral axon that innervates the epidermis and a central axon that connects to downstream circuits in the spinal cord (Palanca et al., 2013; Katz et al., 2021). Unlike in mammals, fish can often regenerate axons in the CNS (Rasmussen and Sagasti, 2017). To visualize RB neurons, we used the Islet1(ss) enhancer, which drives expression in all touch-sensing neurons (Higashijima et al., 2000; Sagasti et al., 2005). To identify a time point for regeneration experiments, we characterized the structure of tail-innervating RB neuron arbors as development progressed, with the aim of finding a stage when RB neurons were morphologically stable but still expressed the transgene strongly. These analyses led us to choose 48 hpf for axotomies, as approximately two-thirds of RB peripheral arbors in the tail had attained a stable branching pattern (i.e., new branches were no longer added) at this time point, although they continued to expand via scaling growth.

To test if dlk or lzk are required for the regeneration of zebrafish RB central axons, we transiently labeled isolated RB neurons in the tail, laser severed their ascending central axons 200 μm from the cell body, and measured the total length of regenerated axons at 24 hpa (Fig. 5A,B). We were careful to use the minimal effective laser power for axotomy because we observed that even moderate damage and scarring in the spinal cord impedes regeneration. Surprisingly, RB central axons in dlkla231, lzkla232, and dlkla231 lzkla232 double mutants regenerated similar to wt neurons (Fig. 5B). In fact, some central axons regenerated more on average than wt axons, although this effect did not reach significance. These data indicate that unlike zebrafish motor axons, or central axons of sensory neurons in invertebrates and mice, central axons of RB neurons in zebrafish larvae do not require dlk or lzk for regeneration.

Figure 5.
Figure 5.

RB central axons regenerate in dlkla231 lzkla232 mutants. A, Top, Forty-eight hours postfertilization RB axon immediately after axotomy. Magenta highlights the separated distal stump that will degenerate. Bottom, Same neuron 24 h postaxotomy. Blue highlights the regenerated axon. B, Overlaid box and dot plots showing central axon length regenerated in the indicated genotypes. C, Overlaid box and dot plots showing growth of peripheral arbors following central axotomy in the indicated genotypes (see above, Materials and Methods; Table 5 for details of statistical analyses). Scale bar, 100 μm.

To test if dlk or lzk are required for the regeneration of RB peripheral axons, we labeled isolated tail-innervating RB neurons, removed the entire peripheral arbor by severing it at the first peripheral branch point at 48 hpf, and measured regeneration at 24 hpa (Fig. 6A). (When a neuron had two arbors separately innervating the skin, both were removed). Although a few axons in each genotype failed to regrow, most severed peripheral axons in single and double mutants regenerated comparable to wt, whether measured as total axon growth or as percentage regeneration of the severed arbor (Fig. 6B,C). Although all genotypes regenerated on average similar size arbors (Fig. 6B), there were some differences in regeneration between different mutant genotypes when measured as percentage regeneration of the original arbor (Fig. 6C), reflecting that each group had different size arbors to begin with. Thus, neither central nor peripheral RB axons require DLK or LZK for regeneration.

Figure 6.
Figure 6.

RB peripheral axon arbors regenerate in dlkla231 lzkla232 mutants. A, Top, Forty-eight hours postfertilization RB peripheral axon immediately after axotomy of the indicated genotypes. Magenta highlights the separated distal stump that will degenerate. Bottom, Same neurons 24 h postaxotomy. Blue highlights the regenerated axon. B, C, Dot plots showing total regenerated length in each genotype (B) and the percentage of the original axon length regenerated (C). Bars indicate the mean (see above, Materials and Methods; Table 5 for details of statistical analyses). Scale bar, 100 μm.

dlk and lzk are not required for DRG neuron peripheral axon regeneration in adult zebrafish

The finding that DLK and LZK are not required for regeneration of RB axons was surprising as MNs in the same genetic background failed to regenerate, and DLK signaling has been implicated in regeneration of another sensory neuron type, DRG neurons in culture and adult mice (Itoh et al., 2009; Shin et al., 2012). We therefore considered the possibility that RB neurons, which are replaced by DRG neurons over the course of development (Reyes et al., 2004; Rasmussen et al., 2018), might use a repair strategy different from neurons that persist into adulthood. To determine if DRG neurons require DLK and LZK for regeneration, we imaged them in a line stably expressing mCherry in sensory neurons (P2rx3a:LexA,4xLexAop:mCherryla207), in wild-type, dlkla231, lzkla232, and dlkla231 lzkla232 double mutant animals. Some genotypes were crossed to the casper mutant background, which lacks pigmentation (White et al., 2008) to facilitate imaging (see above, Materials and Methods for details of transgenic genotypes). We first examined regeneration of adult (8–11 months old) DRG neurons by severing sensory nerves immediately above and under a scale (Fig. 7), as these nerves were easily accessible for injury with a pulsed ultraviolet laser (Rasmussen et al., 2018). Adult fish were intubated during injury and subsequent imaging sessions (Shorey et al., 2021). By 24 h after axotomy, neurites distal to the cut site had degenerated, and by 96 hpa, scales were again fully innervated, comparable to preaxotomy conditions, in wt, dlkla231, lzkla232, and dlkla231 lzkla232 double mutants (n = 10 wt, n = 7 dlkla231, n = 5 lzkla232, n = 5 double mutants; Fig. 7A,B).

Figure 7.
Figure 7.

DRG axons innervating the adult scale regenerate in dlkla231 lzkla232 mutants. A, Images of DRG neurons innervating the epidermis above scales in adult (8–11 months old) zebrafish in indicated genotypes at 0 h postaxotomy. Top, Immediately after axotomy; bottom, immediately before axotomy. Lightning bolts indicate axotomy sites of individual nerves growing into scales. Nerves innervating anterior scales (orange) are above the scale, whereas nerves innervating posterior scale (blue) are below the anterior scale. Most axons innervating these scales degenerated by 24 hpa (middle column) and reinnervated scales by 96 hpa (right columns). Scales were reinnervated in all wt (n = 10), dlkla231 lzkla232 (n = 5), dlkla231(n = 7, data not shown), and lzkla232 (n = 5, data not shown) mutants. B, Quantification of wt (n = 10) and dlkla231 lzkla232 (n = 5) mutant regeneration 96 hpa (see above, Materials and Methods; Table 5 for details of statistical analyses). Scale bar: 200 μm.

A previous study showing reduced regeneration of DRG peripheral axons in DLK mutant neurons monitored regrowth after injury closer to the cell body (Shin et al., 2012), so we considered the possibility that surface damage of sensory axons might be so common that it does not require DLK/LZK signaling. We thus severed DRG nerves close to the ganglion, using two-photon laser surgery in 4- to 5-week-old juvenile fish (Fig. 8A). At this age, DRG sensory arbors on the scale surface have not yet achieved a fully mature morphology, but tail-innervating neurites were indistinguishable from those of adults. We specifically cut the peripheral nerve exiting the most caudal tail-innervating DRG (potentially the equivalent of the sacral ganglion nerve) at ∼100 μm from the ganglion, resulting in the complete loss of tail fin innervation along the dorsal-most fin rays (Fig. 8B). Surprisingly, the nerve regenerated robustly in both wt and dlkla231 lzkla232 double mutants (n = 5 wt; n = 6 double mutants); by 96 hpa, regenerated axons had reached the end of the tail (Fig. 8C,D). Thus, unlike motor axons, sensory axons of RB and DRG neurons do not require LZK and DLK for regeneration.

Figure 8.
Figure 8.

DRG axons innervating the juvenile tail regenerate in wild-type and dlkla231 lzkla232 mutants. A, Top left, Diagram of juvenile casper fish, showing tail-innervating DRGs in P2rx3a: LexA; 4xLexAop:mCherryla207transgenic fish. Lightning bolt indicates axotomy site. Blue arrow indicates the spinal cord. Legend refers to the image on the right. B, Overview showing the position of DRG cell bodies, sensory nerve layout, and an example of axotomy location in a 4- to 5-week-old dlkla231 lzkla232 mutant zebrafish. Inset, Axotomy site of the caudal-most DRG peripheral nerve. Green highlights separated arbors of the severed DRG nerve, which will degenerate after axotomy. C, Top left, Mutant dlkla231 lzkla232 showing regrowth of the severed nerve 24 h postaxotomy. Bottom left: Tail fin of the same animal showing that the regenerating axons have reached the fin tip. C, Top right, The wt fish showing regrowth of the severed nerve 24 hpa. Bottom right, Tail fin of the same animal showing that the regenerating axons have reached the fin tip. D, Quantification of complete regeneration at 96 h postaxotomy. Axons regenerated in all wt (n = 5) and all dlkla231 lzkla232 mutants (n = 6; see above, Materials and Methods; Table 5 for details of statistical analyses). Scale bars: 100 μm.

Spared branches of damaged RB peripheral axons sprout excessively in dlkla231 lzkla232 double mutants

DLK and LZK contribute to injury-induced sprouting of corticospinal neurons in mice (Saikia et al., 2022). Axon regeneration from severed neurites, regenerative sprouting from spared branches of damaged axons, and collateral sprouting from undamaged axons are distinct processes (Steward et al., 2003; Tuszynski and Steward, 2012). Our single RB neuron labeling technique provided an opportunity to ask if DLK or LZK play distinct roles in these processes. To compare the behavior of severed and spared branches of a damaged axon, we severed RB peripheral axons at the second branch point, sparing the first branch (Fig. 9A,B). Surprisingly, partially axotomized peripheral axons in dlkla231 and lzkla232 mutants regenerated more than wt axons; in other words, mutant axons gained more total peripheral axon length than wt axons after axotomy (Fig. 9C). By separately measuring growth of the spared and axotomized branches, we found that regenerative sprouting contributed virtually all of the excess growth, rather than growth from the injury site (Fig. 9D,E). The dlkla231 mutant phenotype was stronger than the lzkla232 mutant phenotype, but excess growth was more pronounced in dlkla231 lzkla232 double mutants than in either single mutant. These results indicate that dlk and lzk inhibit peripheral axon growth specifically in spared branches of damaged axons. To test if this effect is compartment specific, we measured the growth of peripheral axons after severing the central axon of the same neuron. Cutting central axons did not preferentially promote growth of spared peripheral axons in mutants, indicating that growth inhibition by DLK and LZK occurs locally, within a cellular compartment (Fig. 5C).

Figure 9.
Figure 9.

Spared arbors of damaged RB neurons sprout excessively in dlkla231 lzkla232 mutants. A, Diagram of partial RB peripheral axotomy assay, which differentiates between regeneration from the cut site and regenerative sprouting from spared branches. B, Top, Forty-eight hours postfertilization RB axons immediately after axotomy in the indicated genotypes. Lightning bolt indicates axotomy site. Magenta highlights the separated distal stump that will degenerate. Far right, A neuron expressing rescue cDNA; expression of the rescue transgene in the cell body is shown below. Bottom, Same neurons 24 h postaxotomy. Blue highlights regenerated axons; orange highlights the spared branch. C–E, Box and dot plots showing total new growth, including both from the axotomy site and spared branch (C), percentage regeneration from just the injury site (D), and percentage increase of the spared branch (E; see above, Materials and Methods; Table 5 for details of statistical analyses). Scale bar: 100 μm.

To test if DLK acts cell autonomously to inhibit regenerative sprouting, we expressed dlk cDNA in dlkla231 mutants. Like with MNs, strong overexpression of dlk cDNA with the Gal4/UAS system was toxic to RB neurons, causing cell death by 24 hpf (data not shown). As a result, we expressed lower levels of the cDNA (Crest3:DLK-T2A-GFP, coinjected with Isl1[SS]:Gal4 and UAS:DsRed) in RB neurons. Expression of the dlk cDNA modestly reduced axon sprouting when measured as a percentage of the initial arbor size regenerated. This effect did not reach significance when compared with the dlkla231 mutants (Fig. 9C–E), but the rescued single mutants were significantly different from dlkla231 lzkla232 double mutants. However, when measured as total axon length regenerated, there was no apparent rescue with dlk cDNA. These ambiguous results may reflect the difficulty of achieving the appropriate level of cDNA expression for rescue but could also suggest that these proteins do not act strictly cell autonomously to limit regenerative sprouting of spared branches in RB neurons.

dlk and lzk are general inhibitors of collateral sprouting in sensory neurons

Given our unexpected finding that spared branches of damaged RB neurons grew more in mutants than wt animals, we wondered if increased axon growth in dlkla231 and lzkla232 mutants was specific to injured axons, or if sensory axon growth was generally disinhibited in these mutants. As axon growth during development is limited by repulsive tiling interactions (Sagasti et al., 2005; Grueber and Sagasti, 2010), increased growth potential in dlkla231 lzkla232 mutants could be masked by tiling. We therefore compared peripheral axon growth in mutants and wt animals in which we relieved axon tiling by ablating an entire trigeminal ganglion, as larval trigeminal sensory neurons are similar to larval RB neurons (Gau et al., 2013; Palanca et al., 2013). Ablating a trigeminal ganglion early in development (24 hpf) allows axons from the remaining ganglion to grow into the denervated side of the head significantly more than in nonablated control animals because axon arbors are not limited by their contralateral counterparts (Sagasti et al., 2005). However, this growth ability diminishes by 78 hpf (O'Brien et al., 2009b). To reduce variegation of the reporter (i.e., silencing in some cells), we selected homozygous transgenic embryos (Isl1[SS]:Gal4;UAS:DsRed) in which most, if not all, trigeminal cell bodies were labeled. Although axons from each ganglion do not stop abruptly at a sharp midline, they rarely reach the contralateral eye in wild-type animals. Twenty-four hours after ganglion ablation, axons of the spared ganglion grew markedly farther into the denervated side of the head in dlkla231 lzkla232 than in wt animals (Fig. 10A). Measuring the total axon length that grew over the contralateral eye revealed that dlk and lzk inhibit collateral axon sprouting of trigeminal neurons, even in uninjured neurons (Fig. 10B).

Figure 10.
Figure 10.

Trigeminal axons grow excessively in dlkla231 lzkla232 mutants after ablation of the contralateral ganglion. A, Images and diagram of trigeminal axons in zebrafish heads at 78 hpf. Confocal images show two separate examples of wt (left) and dlkla231 lzkla232 mutant (right) fish. Insets, Magnification of a region over the eye, with axons traced in orange. Bottom, Diagram depictions of the result. B, Dot plot showing total axon length that grew over the contralateral eye. Error bar indicates the mean (see above, Materials and Methods; Table 5 for details of statistical analyses). Scale bar, 500 μm.

Discussion

This study establishes and characterizes zebrafish mutants in the dlk and lzk genes, complementing existing worm, fly, and mouse models to study the function of these critical neuronal injury regulators (Jin and Zheng, 2019). Zebrafish offer powerful advantages over other vertebrate models, including the ability to label single neurons of different types and precisely injure them with laser axotomy, thus making it possible to distinguish the responses of different neurite branches to injury. Using this approach, we found that dlk and lzk are required cell autonomously and, partially redundantly, for MN regeneration in larval zebrafish. By contrast, dlk and lzk are not required for axon regeneration in larval RB touch-sensing neurons or adult DRG neurons. However, these kinases negatively regulate the sprouting of spared sensory neuron peripheral arbors, both within the injured neuron and in uninjured neighboring neurons. These findings reveal cell-type-specific dlk and lzk functions and highlight the mechanistic differences between different kinds of regenerative growth, which can be promoted or inhibited by the same signaling molecules.

Invertebrates only have one DLK-related gene, but the existence of a close relative of DLK, namely, LZK, in vertebrates raises the possibility that these MAP3Ks act redundantly (Jin and Zheng, 2019). Despite their similarity, however, DLK lacks a key calcium-binding domain found in LZK and in invertebrate DLK homologs, suggesting that these kinases may be activated in different ways. Studies using optic nerve crush and traumatic brain injury models found that inhibition of both dlk and lzk offered the strongest protection from cell death (Welsbie et al., 2019), indicating that they play overlapping roles in promoting injury-induced death. Similarly, DLK and LZK are required redundantly for PTEN-deletion-induced regeneration of mouse corticospinal neurons and injury-induced corticospinal neuron sprouting (Saikia et al., 2022). By contrast, other studies suggest distinct, cell-type-specific gene functions, including the roles of LZK in activating astrocytes (Chen et al., 2018). We directly addressed the potential for redundancy between DLK and LZK in zebrafish axon regeneration by comparing each mutant with double mutants. In motor axons, dlkla231 mutants had a modest regeneration defect, but dlkla231 lzkla232 double mutants had a strong defect, suggesting partial redundancy. As dlkla231 has an in-frame deletion of the kinase domain, it is unlikely to trigger genetic compensation (Rossi et al., 2015; El-Brolosy et al., 2019; Ma et al., 2019). The lzkla232 mutants, however, have a premature stop codon, which could trigger nonsense-mediated RNA decay and thus genetic compensation, perhaps explaining why lzkla232 mutants did not have a strong motor axon regeneration defect on their own. The fact that a few double mutant motor axons were able to regenerate may suggest compensatory contributions from other MAP3Ks in the MLK family or parallel proregenerative pathways. We found a similar pattern for the suppression of RB neuron regenerative sprouting (sprouting was increased in dlkla231 animals, but it was more pronounced in dlkla231 lzkla232 double mutants) and only saw motor dendrite overgrowth in double mutants. Together, these results suggest a partially redundant or compensatory relationship between these two kinases in axon regeneration, similar to their roles in the regeneration of corticospinal neurons in PTEN mutant mice (Saikia et al., 2022).

DLK promotes axon regeneration in many different types of neurons and organisms, but in larval Drosophila sensory neurons it is dispensable for dendrite regeneration, although it is required for axon regeneration in the same cells (Stone et al., 2014). This observation indicates that axon and dendrite regeneration are mechanistically distinct, and indeed other factors differentially affect these processes (Nye et al., 2020). Vertebrate sensory neurons, although similar in many respects to Drosophila counterparts, have sensory axons rather than dendrites, as defined by physiological features and cytoskeletal organization (Shorey et al., 2021), so we initially hypothesized that DLK and LZK would be required for regeneration of both central and peripheral sensory axons. Surprisingly, however, central RB, peripheral RB, and peripheral DRG axons regenerated normally in dlkla231 lzkla232 mutants. These findings suggest that another, yet unknown, pathway is used by fish somatosensory neurons to detect axon damage and activate the axon regeneration program. The intriguing observation that motor neuron dendrites overgrow in dlkla231 lzkla232 mutants following axotomy and failure of axon regeneration suggests a potential antagonistic interplay between axon and dendrite regeneration, as has been noted in CNS neurons (Beckers et al., 2019; Beckers and Moons, 2019).

When an entire nerve is damaged, it is difficult to deconvolute axon regeneration from a severed axon stump, regenerative sprouting from spared branches, and collateral sprouting from neighboring neurons (Steward et al., 2003; Tuszynski and Steward, 2012). The balance of these types of growth, however, has important functional consequences, particularly for sensory neurons. For example, RB sensory axons in larval zebrafish tile to innervate discrete, minimally overlapping territories in the skin that provide spatial information necessary for appropriate behavioral responses to touch (Sagasti et al., 2005). The balance between true regeneration and collateral sprouting thus determines the sensory map of the periphery. Our single-cell labeling method allowed us to directly address this issue, revealing that DLK, potentially with some contribution from LZK, is required to limit sprouting from uninjured axon branches, although it did not affect regeneration from severed axon branches. This finding emphasizes that forming a growth cone in a damaged axon branch, which has experienced cytoskeletal disruption, calcium influx, and local mitochondrial dysfunction, is a distinct process from reactivating growth in a dormant, uninjured axon branch. Because it is critical for sensory neurons to restore their spatial sensory map in the periphery, limiting sprouting may be as functionally important as promoting new growth from an injured branch. Indeed, excessive neuronal sprouting induced by pathologic conditions such as atopic dermatitis, or selective serotonin reuptake inhibitors treatment, is associated with oversensitivity and itch in the skin (Han and Dong, 2014; Tominaga and Takamori, 2014; Morita et al., 2015).

Our findings that DLK and LZK limit injury-induced axon sprouting in RB and trigeminal neurons contrast with a recent report that DLK and LZK promote injury-induced axon sprouting of mouse corticospinal neurons in both wild-type and PTEN-deleted animals (Saikia et al., 2022). Similar to our trigeminal experiment, sprouting axons in those mouse experiments were from uninjured contralateral neurons. However, the sprouting neurons in our experiments were different in several fundamental ways (e.g., zebrafish vs mouse, larvae vs adult, PNS vs CNS), indicating that DLK and LZK have evolved to promote different injury responses in different cells. Collateral sprouting benefits recovery in some instances by allowing compensatory circuits to form and restore function. However, collateral sprouting of cutaneous sensory axons can be detrimental as sprouting alters the peripheral sensory map, potentially creating mismatches between sensory territories and central circuits that result in inappropriate behavioral responses, such as “escaping” toward, rather than away from, a threatening stimulus (Sagasti et al., 2005). Thus, injury response pathways have evolved to activate distinct responses appropriate to the function of different cell types.

The cellular site in which DLK acts to limit sprouting remains unclear in our experiments because expressing DLK cDNA only modestly reduced sprouting in dlkla231 RB neurons. Achieving rescue experimentally may be difficult, as overexpressing DLK was toxic to neurons, suggesting that precise regulation of DLK levels is required for an optimal injury response. These findings also allow the intriguing possibility that DLK may act non-cell autonomously to limit sprouting. For example, DLK could function in the epidermal cells innervated by RB axons as DLK regulates epidermal differentiation and integrity (Robitaille et al., 2005; Simard-Bisson et al., 2017), or in immune cells activated by the injury as DLK and LZK regulate microglial and astrocyte responses to injury in the CNS (Chen et al., 2018; Wlaschin et al., 2018). Wherever DLK is functioning to limit sprouting, our findings reveal that regeneration from an injured growth cone and sprouting from uninjured axon branches are mechanistically distinct processes that differentially require DLK signaling.

Our discovery that zebrafish DLK and LZK are required to promote axon growth in one context (motor axon regeneration), and restrain it in others (growth of RB spared branches and trigeminal neurons) echoes the context-specific dual roles for DLK homologs in other neurons. For example, in addition to its differential effects on axon and dendrite regeneration in Drosophila sensory neurons (Stone et al., 2014), excess DLK promotes axon growth and inhibits dendrite growth in the same neurons (Wang et al., 2013). Moreover, although DLK is required for axon regeneration in Drosophila sensory neurons, it also promotes an opposing neuroprotective response that inhibits axon regeneration (L. Chen et al., 2016). Thus, DLK has the potential to both activate and inhibit regeneration in these neurons, depending on the balance of its outputs. Similarly, in C. elegans sensory neurites, excess DLK resulting from loss of a negative regulator can inhibit axon regeneration (Yan et al., 2009) or promote developmental overgrowth (Zheng et al., 2020), depending on which regulator was reduced.

Our study adds to a growing list of examples of distinct DLK functions in different cell types and conditions (Jin and Zheng, 2019). These diverse outcomes could be explained by the permissiveness versus nonpermissiveness of the environment, expression levels of these signaling proteins, subcellular localization, the mode of activation, or even the duration of the signal, all of which could lead to the assembly of different signaling complexes that activate different responses (Goodwani et al., 2020). Understanding these diverse molecular processes, categorizing potential outcomes, and determining the neuronal cell types in which they occur in vivo are prerequisites to considering dlk and lzk as targets in the treatment of axonal neuropathies and traumatic injuries.

Footnotes

  • This study was supported by National Institute of Neurological Disorders and Stroke Grant R21NS090027 (M.M.R. and A.S.) and National Institute of Arthritis and Musculoskeletal and Skin Diseases Grant R01AR064582 (A.S.). K.P.A. was supported by a University of California, Los Angeles Cota-Robles Fellowship, National Science Foundation Bridge to the Doctorate Fellowship, and National Institute of Neurological Disorders and Stroke F31 Predoctoral Fellowship Grant NS106742. We thank members of the Sagasti Lab for comments on the manuscript, Son Giang and Yuan Dong for fish care, and Adam Langenbacher for help with data presentation.

  • The authors declare no competing financial interests.

  • Correspondence should be addressed to Alvaro Sagasti at sagasti{at}mcdb.ucla.edu

References

  1. ↵
    2. Beckers A,
    3. Moons L
    (2019) Dendritic shrinkage after injury: a cellular killer or a necessity for axonal regeneration? Neural Regen Res 14:1313–1316. doi:10.4103/1673-5374.253505 pmid:30860164
    OpenUrlCrossRefPubMed
  2. ↵
    2. Beckers A,
    3. Van Dyck A,
    4. Bollaerts I,
    5. Van Houcke J,
    6. Lefevere E,
    7. Andries L,
    8. Agostinone J,
    9. Van Hove I,
    10. Di Polo A,
    11. Lemmens K,
    12. Moons L
    (2019) An antagonistic axon-dendrite interplay enables efficient neuronal repair in the adult zebrafish central nervous system. Mol Neurobiol 56:3175–3192. doi:10.1007/s12035-018-1292-5 pmid:30105671
    OpenUrlCrossRefPubMed
  3. ↵
    2. Chen L,
    3. Nye DM,
    4. Stone MC,
    5. Weiner AT,
    6. Gheres KW,
    7. Xiong X,
    8. Collins CA,
    9. Rolls MM
    (2016) Mitochondria and caspases tune Nmnat-mediated stabilization to promote axon regeneration. PLoS Genet 12:e1006503. doi:10.1371/journal.pgen.1006503 pmid:27923046
    OpenUrlCrossRefPubMed
  4. ↵
    2. Chen M,
    3. Geoffroy CG,
    4. Wong HN,
    5. Tress O,
    6. Nguyen MT,
    7. Holzman LB,
    8. Jin Y,
    9. Zheng B
    (2016) Leucine zipper-bearing kinase promotes axon growth in mammalian central nervous system neurons. Sci Rep 6:31482. doi:10.1038/srep31482 pmid:27511108
    OpenUrlCrossRefPubMed
  5. ↵
    2. Chen M,
    3. Geoffroy CG,
    4. Meves JM,
    5. Narang A,
    6. Li Y,
    7. Nguyen MT,
    8. Khai VS,
    9. Kong X,
    10. Steinke CL,
    11. Carolino KI,
    12. Elzière L,
    13. Goldberg MP,
    14. Jin Y,
    15. Zheng B
    (2018) Leucine zipper-bearing kinase is a critical regulator of astrocyte reactivity in the adult mammalian CNS. Cell Rep 22:3587–3597. doi:10.1016/j.celrep.2018.02.102 pmid:29590625
    OpenUrlCrossRefPubMed
  6. ↵
    2. Craig EA,
    3. Stevens MV,
    4. Vaillancourt RR,
    5. Camenisch TD
    (2008) MAP3Ks as central regulators of cell fate during development. Dev Dyn 237:3102–3114. doi:10.1002/dvdy.21750 pmid:18855897
    OpenUrlCrossRefPubMed
  7. ↵
    2. Curcio M,
    3. Bradke F
    (2018) Axon regeneration in the central nervous system: facing the challenges from the inside. Annu Rev Cell Dev Biol 34:495–521. doi:10.1146/annurev-cellbio-100617-062508 pmid:30044649
    OpenUrlCrossRefPubMed
  8. ↵
    2. Dickson HM,
    3. Zurawski J,
    4. Zhang H,
    5. Turner DL,
    6. Vojtek AB
    (2010) POSH is an intracellular signal transducer for the axon outgrowth inhibitor Nogo66. J Neurosci 30:13319–13325. doi:10.1523/JNEUROSCI.1324-10.2010 pmid:20926658
    OpenUrlAbstract/FREE Full Text
  9. ↵
    2. El-Brolosy MA,
    3. Kontarakis Z,
    4. Rossi A,
    5. Kuenne C,
    6. Günther S,
    7. Fukuda N,
    8. Kikhi K,
    9. Boezio GLM,
    10. Takacs CM,
    11. Lai S-L,
    12. Fukuda R,
    13. Gerri C,
    14. Giraldez AJ,
    15. Stainier DYR
    (2019) Genetic compensation triggered by mutant mRNA degradation. Nature 568:193–197. doi:10.1038/s41586-019-1064-z
    OpenUrlCrossRefPubMed
  10. ↵
    2. Gallo KA,
    3. Johnson GL
    (2002) Mixed-lineage kinase control of JNK and p38 MAPK pathways. Nat Rev Mol Cell Biol 3:663–672. doi:10.1038/nrm906 pmid:12209126
    OpenUrlCrossRefPubMed
  11. ↵
    2. Gau P,
    3. Poon J,
    4. Ufret-Vincenty C,
    5. Snelson CD,
    6. Gordon SE,
    7. Raible DW,
    8. Dhaka A
    (2013) The zebrafish ortholog of TRPV1 is required for heat-induced locomotion. J Neurosci 33:5249–5260. doi:10.1523/JNEUROSCI.5403-12.2013 pmid:23516290
    OpenUrlAbstract/FREE Full Text
  12. ↵
    2. Ghosh AS,
    3. Wang B,
    4. Pozniak CD,
    5. Chen M,
    6. Watts RJ,
    7. Lewcock JW
    (2011) DLK induces developmental neuronal degeneration via selective regulation of proapoptotic JNK activity. J Cell Biol 194:751–764. doi:10.1083/jcb.201103153 pmid:21893599
    OpenUrlAbstract/FREE Full Text
  13. ↵
    2. Goodwani S,
    3. Fernandez C,
    4. Acton PJ,
    5. Buggia-Prevot V,
    6. McReynolds ML,
    7. Ma J,
    8. Hu CH,
    9. Hamby ME,
    10. Jiang Y,
    11. Le K,
    12. Soth MJ,
    13. Jones P,
    14. Ray WJ
    (2020) Dual leucine zipper kinase is constitutively active in the adult mouse brain and has both stress-induced and homeostatic functions. Int J Mol Sci 21:4849. doi:10.3390/ijms21144849
    OpenUrlCrossRefPubMed
  14. ↵
    2. Grueber WB,
    3. Sagasti A
    (2010) Self-avoidance and tiling: mechanisms of dendrite and axon spacing. Cold Spring Harb Perspect Biol 2:a001750. doi:10.1101/cshperspect.a001750 pmid:20573716
    OpenUrlAbstract/FREE Full Text
  15. ↵
    2. Hammarlund M,
    3. Nix P,
    4. Hauth L,
    5. Jorgensen EM,
    6. Bastiani M
    (2009) Axon regeneration requires a conserved MAP kinase pathway. Science 323:802–806. doi:10.1126/science.1165527 pmid:19164707
    OpenUrlAbstract/FREE Full Text
  16. ↵
    2. Han L,
    3. Dong X
    (2014) Itch mechanisms and circuits. Annu Rev Biophys 43:331–355. doi:10.1146/annurev-biophys-051013-022826 pmid:24819620
    OpenUrlCrossRefPubMed
  17. ↵
    2. Higashijima S,
    3. Hotta Y,
    4. Okamoto H
    (2000) Visualization of cranial motor neurons in live transgenic zebrafish expressing green fluorescent protein under the control of the islet-1 promoter/enhancer. J Neurosci 20:206–218. pmid:10627598
    OpenUrlAbstract/FREE Full Text
  18. ↵
    2. Hirai S-I,
    3. Cui DF,
    4. Miyata T,
    5. Ogawa M,
    6. Kiyonari H,
    7. Suda Y,
    8. Aizawa S,
    9. Banba Y,
    10. Ohno S
    (2006) The c-Jun N-terminal kinase activator dual leucine zipper kinase regulates axon growth and neuronal migration in the developing cerebral cortex. J Neurosci 26:11992–12002. doi:10.1523/JNEUROSCI.2272-06.2006 pmid:17108173
    OpenUrlAbstract/FREE Full Text
  19. ↵
    2. Hirai S-I,
    3. Banba Y,
    4. Satake T,
    5. Ohno S
    (2011) Axon formation in neocortical neurons depends on stage-specific regulation of microtubule stability by the dual leucine zipper kinase-c-Jun N-terminal kinase pathway. J Neurosci 31:6468–6480. doi:10.1523/JNEUROSCI.5038-10.2011 pmid:21525288
    OpenUrlAbstract/FREE Full Text
  20. ↵
    2. Issa FA,
    3. Mazzochi C,
    4. Mock AF,
    5. Papazian DM
    (2011) Spinocerebellar ataxia type 13 mutant potassium channel alters neuronal excitability and causes locomotor deficits in zebrafish. J Neurosci 31:6831–6841. doi:10.1523/JNEUROSCI.6572-10.2011 pmid:21543613
    OpenUrlAbstract/FREE Full Text
  21. ↵
    2. Itoh A,
    3. Horiuchi M,
    4. Bannerman P,
    5. Pleasure D,
    6. Itoh T
    (2009) Impaired regenerative response of primary sensory neurons in ZPK/DLK gene-trap mice. Biochem Biophys Res Commun 383:258–262. doi:10.1016/j.bbrc.2009.04.009 pmid:19358824
    OpenUrlCrossRefPubMed
  22. ↵
    2. Jin Y,
    3. Zheng B
    (2019) Multitasking: dual leucine zipper-bearing kinases in neuronal development and stress management. Annu Rev Cell Dev Biol 35:501–521. doi:10.1146/annurev-cellbio-100617-062644 pmid:31590586
    OpenUrlCrossRefPubMed
  23. ↵
    2. Katz HR,
    3. Menelaou E,
    4. Hale ME
    (2021) Morphological and physiological properties of Rohon–Beard neurons along the zebrafish spinal cord. J Comp Neurol 529:1499–1515. doi:10.1002/cne.25033
    OpenUrlCrossRef
  24. ↵
    2. Kwan KM,
    3. Fujimoto E,
    4. Grabher C,
    5. Mangum BD,
    6. Hardy ME,
    7. Campbell DS,
    8. Parant JM,
    9. Yost HJ,
    10. Kanki JP,
    11. Chien C-B
    (2007) The Tol2kit: a multisite gateway-based construction kit for Tol2 transposon transgenesis constructs. Dev Dyn 236:3088–3099. doi:10.1002/dvdy.21343 pmid:17937395
    OpenUrlCrossRefPubMed
  25. ↵
    2. Letunic I,
    3. Bork P
    (2007) Interactive Tree of Life (iTOL): an online tool for phylogenetic tree display and annotation. Bioinformatics 23:127–128. doi:10.1093/bioinformatics/btl529 pmid:17050570
    OpenUrlCrossRefPubMed
  26. ↵
    2. Li Y,
    3. Ritchie EM,
    4. Steinke CL,
    5. Qi C,
    6. Chen L,
    7. Zheng B,
    8. Jin Y
    (2021) Activation of MAP3K DLK and LZK in Purkinje cells causes rapid and slow degeneration depending on signaling strength. Elife 10:e63509. doi:10.7554/eLife.63509
    OpenUrlCrossRef
  27. ↵
    2. Ma Z,
    3. Zhu P,
    4. Shi H,
    5. Guo L,
    6. Zhang Q,
    7. Chen Y,
    8. Chen S,
    9. Zhang Z,
    10. Peng J,
    11. Chen J
    (2019) PTC-bearing mRNA elicits a genetic compensation response via Upf3a and COMPASS components. Nature 568:259–263. doi:10.1038/s41586-019-1057-y pmid:30944473
    OpenUrlCrossRefPubMed
  28. ↵
    2. Miller BR,
    3. Press C,
    4. Daniels RW,
    5. Sasaki Y,
    6. Milbrandt J,
    7. DiAntonio A
    (2009) A dual leucine kinase-dependent axon self-destruction program promotes Wallerian degeneration. Nat Neurosci 12:387–389. doi:10.1038/nn.2290 pmid:19287387
    OpenUrlCrossRefPubMed
  29. ↵
    2. Miller MA,
    3. Pfeiffer W,
    4. Schwartz T
    (2010) Creating the CIPRES Science Gateway for inference of large phylogenetic trees. Paper presented at Gateway Computing Environments Workshop, New Orleans, November.
  30. ↵
    2. Morita T,
    3. McClain SP,
    4. Batia LM,
    5. Pellegrino M,
    6. Wilson SR,
    7. Kienzler MA,
    8. Lyman K,
    9. Olsen ASB,
    10. Wong JF,
    11. Stucky CL,
    12. Brem RB,
    13. Bautista DM
    (2015) HTR7 mediates serotonergic acute and chronic itch. Neuron 87:124–138. doi:10.1016/j.neuron.2015.05.044 pmid:26074006
    OpenUrlCrossRefPubMed
  31. ↵
    2. Nakata K,
    3. Abrams B,
    4. Grill B,
    5. Goncharov A,
    6. Huang X,
    7. Chisholm AD,
    8. Jin Y
    (2005) Regulation of a DLK-1 and p38 MAP kinase pathway by the ubiquitin ligase RPM-1 is required for presynaptic development. Cell 120:407–420. doi:10.1016/j.cell.2004.12.017 pmid:15707898
    OpenUrlCrossRefPubMed
  32. ↵
    2. Nye DMR,
    3. Albertson RM,
    4. Weiner AT,
    5. Hertzler JI,
    6. Shorey M,
    7. Goberdhan DCI,
    8. Wilson C,
    9. Janes KA,
    10. Rolls MM
    (2020) The receptor tyrosine kinase Ror is required for dendrite regeneration in Drosophila neurons. PLoS Biol 18:e3000657. doi:10.1371/journal.pbio.3000657 pmid:32163406
    OpenUrlCrossRefPubMed
  33. ↵
    2. O'Brien GS,
    3. Rieger S,
    4. Martin SM,
    5. Cavanaugh AM,
    6. Portera-Cailliau C,
    7. Sagasti A
    (2009a) Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos. J Vis Exp 2009:1129.
    OpenUrl
  34. ↵
    2. O'Brien GS,
    3. Martin SM,
    4. Söllner C,
    5. Wright GJ,
    6. Becker CG,
    7. Portera-Cailliau C,
    8. Sagasti A
    (2009b) Developmentally regulated impediments to skin reinnervation by injured peripheral sensory axon terminals. Curr Biol 19:2086–2090. doi:10.1016/j.cub.2009.10.051 pmid:19962310
    OpenUrlCrossRefPubMed
  35. ↵
    2. Palanca AMS,
    3. Lee S-L,
    4. Yee LE,
    5. Joe-Wong C,
    6. Trinh LA,
    7. Hiroyasu E,
    8. Husain M,
    9. Fraser SE,
    10. Pellegrini M,
    11. Sagasti A
    (2013) New transgenic reporters identify somatosensory neuron subtypes in larval zebrafish. Dev Neurobiol 73:152–167. doi:10.1002/dneu.22049 pmid:22865660
    OpenUrlCrossRefPubMed
  36. ↵
    2. Rasmussen JP,
    3. Sagasti A
    (2017) Learning to swim, again: axon regeneration in fish. Exp Neurol 287:318–330. doi:10.1016/j.expneurol.2016.02.022 pmid:26940084
    OpenUrlCrossRefPubMed
  37. ↵
    2. Rasmussen JP,
    3. Vo N-T,
    4. Sagasti A
    (2018) Fish scales dictate the pattern of adult skin innervation and vascularization. Dev Cell 46:344–359.e4. doi:10.1016/j.devcel.2018.06.019 pmid:30032992
    OpenUrlCrossRefPubMed
  38. ↵
    2. Reyes R,
    3. Haendel M,
    4. Grant D,
    5. Melancon E,
    6. Eisen JS
    (2004) Slow degeneration of zebrafish Rohon–Beard neurons during programmed cell death. Dev Dyn 229:30–41. doi:10.1002/dvdy.10488 pmid:14699575
    OpenUrlCrossRefPubMed
  39. ↵
    2. Robitaille H,
    3. Proulx R,
    4. Robitaille K,
    5. Blouin R,
    6. Germain L
    (2005) The mitogen-activated protein kinase kinase kinase dual leucine zipper-bearing kinase (DLK) acts as a key regulator of keratinocyte terminal differentiation. J Biol Chem 280:12732–12741. doi:10.1074/jbc.M411619200 pmid:15695824
    OpenUrlAbstract/FREE Full Text
  40. ↵
    2. Rosenberg AF,
    3. Wolman MA,
    4. Franzini-Armstrong C,
    5. Granato M
    (2012) In vivo nerve-macrophage interactions following peripheral nerve injury. J Neurosci 32:3898–3909. doi:10.1523/JNEUROSCI.5225-11.2012 pmid:22423110
    OpenUrlAbstract/FREE Full Text
  41. ↵
    2. Rossi A,
    3. Kontarakis Z,
    4. Gerri C,
    5. Nolte H,
    6. Hölper S,
    7. Krüger M,
    8. Stainier DYR
    (2015) Genetic compensation induced by deleterious mutations but not gene knockdowns. Nature 524:230–233. doi:10.1038/nature14580 pmid:26168398
    OpenUrlCrossRefPubMed
  42. ↵
    2. Sagasti A,
    3. Guido MR,
    4. Raible DW,
    5. Schier AF
    (2005) Repulsive interactions shape the morphologies and functional arrangement of zebrafish peripheral sensory arbors. Curr Biol 15:804–814. doi:10.1016/j.cub.2005.03.048 pmid:15886097
    OpenUrlCrossRefPubMed
  43. ↵
    2. Saikia JM,
    3. Chavez-Martinez CL,
    4. Kim ND,
    5. Allibhoy S,
    6. Kim HJ,
    7. Simonyan L,
    8. Smadi S,
    9. Tsai KM,
    10. Romaus-Sanjurjo D,
    11. Jin Y,
    12. Zheng B
    (2022) A critical role for DLK and LZK in axonal repair in the mammalian spinal cord. J Neurosci 42:3716–3732. doi:10.1523/JNEUROSCI.2495-21.2022 pmid:35361703
    OpenUrlAbstract/FREE Full Text
    2. Schindelin J,
    3. Arganda-Carreras I,
    4. Frise E,
    5. Kaynig V,
    6. Longair M,
    7. Pietzsch T,
    8. Preibisch S,
    9. Rueden C,
    10. Saalfeld S,
    11. Schmid B,
    12. Tinevez J-Y,
    13. White DJ,
    14. Hartenstein V,
    15. Eliceiri K,
    16. Tomancak P,
    17. Cardona A
    (2012) Fiji: an open-source platform for biological-image analysis. Nat Methods 9:676–682. doi:10.1038/nmeth.2019 pmid:22743772
    OpenUrlCrossRefPubMed
  45. ↵
    2. Shin JE,
    3. Cho Y,
    4. Beirowski B,
    5. Milbrandt J,
    6. Cavalli V,
    7. DiAntonio A
    (2012) Dual leucine zipper kinase is required for retrograde injury signaling and axonal regeneration. Neuron 74:1015–1022. doi:10.1016/j.neuron.2012.04.028 pmid:22726832
    OpenUrlCrossRefPubMed
  46. ↵
    2. Shorey M,
    3. Rao K,
    4. Stone MC,
    5. Mattie FJ,
    6. Sagasti A,
    7. Rolls MM
    (2021) Microtubule organization of vertebrate sensory neurons in vivo. Dev Biol 478:1–12. doi:10.1016/j.ydbio.2021.06.007
    OpenUrlCrossRefPubMed
  47. ↵
    2. Simard-Bisson C,
    3. Bidoggia J,
    4. Larouche D,
    5. Guérin SL,
    6. Blouin R,
    7. Hirai S-I,
    8. Germain L
    (2017) A Role for DLK in microtubule reorganization to the cell periphery and in the maintenance of desmosomal and tight junction integrity. J Invest Dermatol 137:132–141. doi:10.1016/j.jid.2016.07.035 pmid:27519653
    OpenUrlCrossRefPubMed
  48. ↵
    2. Stamatakis A
    (2014) RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30:1312–1313. doi:10.1093/bioinformatics/btu033 pmid:24451623
    OpenUrlCrossRefPubMed
  49. ↵
    2. Steward O,
    3. Zheng B,
    4. Tessier-Lavigne M
    (2003) False resurrections: distinguishing regenerated from spared axons in the injured central nervous system. J Comp Neurol 459:1–8. doi:10.1002/cne.10593 pmid:12629662
    OpenUrlCrossRefPubMed
  50. ↵
    2. Stone MC,
    3. Albertson RM,
    4. Chen L,
    5. Rolls MM
    (2014) Dendrite injury triggers DLK-independent regeneration. Cell Rep 6:247–253. doi:10.1016/j.celrep.2013.12.022 pmid:24412365
    OpenUrlCrossRefPubMed
  51. ↵
    2. Summers DW,
    3. Milbrandt J,
    4. DiAntonio A
    (2018) Palmitoylation enables MAPK-dependent proteostasis of axon survival factors. Proc Natl Acad Sci U S A 115:E8746–E8754. doi:10.1073/pnas.1806933115 pmid:30150401
    OpenUrlAbstract/FREE Full Text
  52. ↵
    2. Talbot JC,
    3. Amacher SL
    (2014) A streamlined CRISPR pipeline to reliably generate zebrafish frameshifting alleles. Zebrafish 11:583–585. doi:10.1089/zeb.2014.1047 pmid:25470533
    OpenUrlCrossRefPubMed
  53. ↵
    2. Tedeschi A,
    3. Bradke F
    (2013) The DLK signalling pathway–a double-edged sword in neural development and regeneration. EMBO Rep 14:605–614. doi:10.1038/embor.2013.64 pmid:23681442
    OpenUrlAbstract/FREE Full Text
  54. ↵
    2. Thisse B,
    3. Heyer V,
    4. Lux A,
    5. Alunni V,
    6. Degrave A,
    7. Seiliez I,
    8. Kirchner J,
    9. Parkhill J-P,
    10. Thisse C
    (2004) Spatial and temporal expression of the zebrafish genome by large-scale in situ hybridization screening. Methods Cell Biol 77:505–519. doi:10.1016/s0091-679x(04)77027-2 pmid:15602929
    OpenUrlCrossRefPubMed
  55. ↵
    2. Tominaga M,
    3. Takamori K
    (2014) Itch and nerve fibers with special reference to atopic dermatitis: therapeutic implications. J Dermatol 41:205–212. doi:10.1111/1346-8138.12317 pmid:24628070
    OpenUrlCrossRefPubMed
  56. ↵
    2. Tuszynski MH,
    3. Steward O
    (2012) Concepts and methods for the study of axonal regeneration in the CNS. Neuron 74:777–791. doi:10.1016/j.neuron.2012.05.006 pmid:22681683
    OpenUrlCrossRefPubMed
  57. ↵
    2. Wang X,
    3. Kim JH,
    4. Bazzi M,
    5. Robinson S,
    6. Collins CA,
    7. Ye B
    (2013) Bimodal control of dendritic and axonal growth by the dual leucine zipper kinase pathway. PLoS Biol 11:e1001572. doi:10.1371/journal.pbio.1001572 pmid:23750116
    OpenUrlCrossRefPubMed
  58. ↵
    2. Watkins TA,
    3. Wang B,
    4. Huntwork-Rodriguez S,
    5. Yang J,
    6. Jiang Z,
    7. Eastham-Anderson J,
    8. Modrusan Z,
    9. Kaminker JS,
    10. Tessier-Lavigne M,
    11. Lewcock JW
    (2013) DLK initiates a transcriptional program that couples apoptotic and regenerative responses to axonal injury. Proc Natl Acad Sci U S A 110:4039–4044. doi:10.1073/pnas.1211074110 pmid:23431164
    OpenUrlAbstract/FREE Full Text
  59. ↵
    2. Welsbie DS, et al
    . (2017) Enhanced functional genomic screening identifies novel mediators of dual leucine zipper kinase-dependent injury signaling in neurons. Neuron 94:1142–1154.e6. doi:10.1016/j.neuron.2017.06.008 pmid:28641113
    OpenUrlCrossRefPubMed
  60. ↵
    2. Welsbie DS,
    3. Ziogas NK,
    4. Xu L,
    5. Kim B-J,
    6. Ge Y,
    7. Patel AK,
    8. Ryu J,
    9. Lehar M,
    10. Alexandris AS,
    11. Stewart N,
    12. Zack DJ,
    13. Koliatsos VE
    (2019) Targeted disruption of dual leucine zipper kinase and leucine zipper kinase promotes neuronal survival in a model of diffuse traumatic brain injury. Mol Neurodegener 14:44.
    OpenUrl
  61. ↵
    2. Westerfield M,
    3. McMurray JV,
    4. Eisen JS
    (1986) Identified motoneurons and their innervation of axial muscles in the zebrafish. J Neurosci 6:2267–2277. pmid:3746409
    OpenUrlAbstract/FREE Full Text
  62. ↵
    2. White RM,
    3. Sessa A,
    4. Burke C,
    5. Bowman T,
    6. LeBlanc J,
    7. Ceol C,
    8. Bourque C,
    9. Dovey M,
    10. Goessling W,
    11. Burns CE,
    12. Zon LI
    (2008) Transparent adult zebrafish as a tool for in vivo transplantation analysis. Cell Stem Cell 2:183–189. doi:10.1016/j.stem.2007.11.002 pmid:18371439
    OpenUrlCrossRefPubMed
  63. ↵
    2. Wlaschin JJ,
    3. Gluski JM,
    4. Nguyen E,
    5. Silberberg H,
    6. Thompson JH,
    7. Chesler AT,
    8. Le Pichon CE
    (2018) Dual leucine zipper kinase is required for mechanical allodynia and microgliosis after nerve injury. Elife 7:e33910. doi:10.7554/eLife.33910
    OpenUrlCrossRef
  64. ↵
    2. Xiong X,
    3. Wang X,
    4. Ewanek R,
    5. Bhat P,
    6. Diantonio A,
    7. Collins CA
    (2010) Protein turnover of the Wallenda/DLK kinase regulates a retrograde response to axonal injury. J Cell Biol 191:211–223. doi:10.1083/jcb.201006039 pmid:20921142
    OpenUrlAbstract/FREE Full Text
  65. ↵
    2. Yan D,
    3. Jin Y
    (2012) Regulation of DLK-1 kinase activity by calcium-mediated dissociation from an inhibitory isoform. Neuron 76:534–548. doi:10.1016/j.neuron.2012.08.043 pmid:23141066
    OpenUrlCrossRefPubMed
  66. ↵
    2. Yan D,
    3. Wu Z,
    4. Chisholm AD,
    5. Jin Y
    (2009) The DLK-1 kinase promotes mRNA stability and local translation in C. elegans synapses and axon regeneration. Cell 138:1005–1018. doi:10.1016/j.cell.2009.06.023 pmid:19737525
    OpenUrlCrossRefPubMed
  67. ↵
    2. Yin C,
    3. Huang G-F,
    4. Sun X-C,
    5. Guo Z,
    6. Zhang JH
    (2017) DLK silencing attenuated neuron apoptosis through JIP3/MA2K7/JNK pathway in early brain injury after SAH in rats. Neurobiol Dis 103:133–143. doi:10.1016/j.nbd.2017.04.006 pmid:28396258
    OpenUrlCrossRefPubMed
  68. ↵
    2. Zheng C,
    3. Atlas E,
    4. Lee HMT,
    5. Jao SLJ,
    6. Nguyen KCQ,
    7. Hall DH,
    8. Chalfie M
    (2020) Opposing effects of an F-box protein and the HSP90 chaperone network on microtubule stability and neurite growth in Caenorhabditis elegans. Development 147:dev189886. doi:10.1242/dev.189886
    OpenUrlAbstract/FREE Full Text
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The MAP3Ks DLK and LZK Direct Diverse Responses to Axon Damage in Zebrafish Peripheral Neurons
Kadidia Pemba Adula, Matthew Shorey, Vasudha Chauhan, Khaled Nassman, Shu-Fan Chen, Melissa M. Rolls, Alvaro Sagasti
Journal of Neuroscience 10 August 2022, 42 (32) 6195-6210; DOI: 10.1523/JNEUROSCI.1395-21.2022
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