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. 2026 Jul 28;86(4):e70054. doi: 10.1002/dneu.70054

Impaired Motor Function in a Zebrafish Stathmin‐2 Knockout Model

Tyler J N Gurberg 1, Ziyaan A Harji 1, Christian J Rampal 1, Joséphine Sacy‐Richer 1, Amy Wang 1, Esteban C Rodríguez 1, Gary A B Armstrong 1,✉
PMCID: PMC13416224  PMID: 42522225

ABSTRACT

Stathmin‐2 (STMN2) is a microtubule‐associated protein that plays a role in the stability of microtubules in axons of the nervous system of animals. In this study, we generated a novel zebrafish STMN2 knockout (KO) model. STMN2 is represented by two genes in the zebrafish genome: stmn2a and stmn2b. Using the CRISPR/Cas9 mutagenic system, we selected founder fish lines harboring frameshift mutations in both genes and bred these together to generate a double stmn2a and stmn2b KO model. Using these models, we observed increased developmental lethality in our double stmn2a and stmn2b KO model and impaired motor function at embryonic stages of development. Examination of the neuromuscular junction (NMJ) revealed a slight increase in the number of orphaned NMJs in trunk musculature as well as a reduction in the amplitude of miniature endplate currents in our double stmn2a and stmn2b KO model. In a final series of experiments, we show impaired ventral root axon regrowth following transection in double stmn2a and stmn2b KO zebrafish. Our findings suggest that while not essential for motor function development, loss of stmn2a and stmn2b expression results in a minor motor phenotype and impairs the ability to regenerate motor axons following injury.

1. Introduction

During vertebrate development, differentiated spinal cord motor neurons extend axons to the periphery, where they form neuromuscular junctions (NMJs) with muscle cells. For proper targeting of muscle cells to occur, growth cones and their axons respond dynamically to extracellular cues by remodeling their microtubule networks. Microtubule networks undergo remodeling through phases of individual microtubule polymerization and depolymerization in a process known as dynamic instability (Mitchison and Kirschner 1984). Microtubule‐associated proteins are a class of proteins involved with microtubule dynamics and include members of the stathmin family of proteins, with humans possessing four members (STMN1–4). The stathmin family of proteins all possess a shared C‐terminal stathmin‐like domain that binds tubulin in a phosphorylation dependent manner by JNK1 (Belmont and Mitchison 1996; Tararuk et al. 2006). Unlike STMN1, STMN2–4 differ in that they are expressed exclusively in the nervous system (Bièche et al. 2003) and possess an N‐terminal motif responsible for vesicular membrane attachment, as well as a targeting signal that directs them to the Golgi complex (Di Paolo et al. 1997; Chauvin and Sobel 2015; Benarroch 2021).

The function of the stathmin family of proteins in motor axon development remains incompletely understood despite both human and mouse motor neurons abundantly expressing STMN2 (Sun et al. 2015; Melamed et al. 2019), suggesting an important role for regulating the cytoskeleton of these cells. STMN2 (also known as SCG10) is upregulated during development and following axonal injury where it is anterogradely trafficked to growth cones (Stein et al. 1988; Shin et al. 2014). Additionally, in individuals with the neurodegenerative disease amyotrophic lateral sclerosis (ALS), STMN2 expression is reduced in motor neurons (Klim et al. 2019; Melamed et al. 2019). STMN2 acts as a microtubule‐destabilizing protein by sequestering soluble tubulin dimers and inducing curvature at microtubule tips, promoting rapid depolymerization and shortening events known as catastrophes (Curmi et al. 1997; Gardner et al. 2013; Gupta et al. 2013). These events are essential for growth cone advancement and responsiveness to guidance cues (Grenningloh et al. 2004; Manna et al. 2007). Investigations examining individual microtubules in vitro, suggest that increasing the molar ratio of STMN2 to tubulin increases the frequency of catastrophe events and microtubule growth rates (Manna et al. 2007). In rat primary cultures of hippocampal cells, siRNA knockdown of Stmn2 expression has been shown to impair neurite growth (Morii et al. 2006) and increase the growth cone surface area (Poulain and Sobel 2010). Similarly, cultured dorsal root ganglion neurons from Stmn2 knockout (KO) mice exhibit impaired axon outgrowth (Krus et al. 2022). These KO mice also display elevated perinatal mortality, and surviving animals develop sensory and motor neuropathy, including NMJ denervation (Krus et al. 2022; Li et al. 2023; López‐Erauskin et al. 2024).

To investigate the role of STMN2 in motor axon development, we generated a zebrafish stmn2 KO model. The zebrafish genome contains two orthologs of STMN2 (stmn2a and stmn2b). Previous wholemount in situ hybridization assays have revealed that both stmn2a and stmn2b are expressed in the nervous system of the zebrafish embryo (Burzynski et al. 2009). Using the CRISPR/Cas9 system, we generated KO models of each gene and bred our fish to generate a double KO model. Using these models, we investigated motor phenotypes. During zebrafish embryogenesis, motor axons rapidly grow and form NMJs in the axial trunk musculature used for swimming behavior. Orchestration of this process must be particularly tuned to effectively establish the circuitry for motor escape responses. In this study, we demonstrate that stmn2a and stmn2b double KO zebrafish display elevated developmental lethality, impaired motor function, a slight increase in the number of orphaned AchRs clusters in muscle cells, and impaired motor axon regrowth following axotomy. Our findings suggest that while not essential for motor axon development, loss of stmn2 expression perturbs the assembly of zebrafish NMJs during development.

2. Methods

2.1. Zebrafish Housing and Maintenance

Adult zebrafish (Danio rerio) of the Tübingen long fin (TL) were maintained according to standard protocols (Westerfield 1995) at 28.5°C and pH 7.5–7.8 under a 14/10 h light/dark cycle in the animal research facility of the Montreal Neurological Institute (MNI) at McGill University (Montreal, Quebec, Canada). Our TL fish line originated from the laboratory of Dr. Pierre Drapeau (Université de Montréal) and later crossed with a TL line maintained by Dr. Edward Ruthazer (McGill University). Eleven generations of our TL fish line have been maintained in our aquatics facility (Aquaneering). Fish were fed three times/day (2× dry food and 1× Artemia). Zebrafish embryos and larvae were raised in system water. Survival assays were conducted by placing 30 five‐day‐old larvae of the same genotype into a single 1.8 L tank housed in our aquatics facility and fed Artemia and dry food daily and raised under the same light/dark cycle as our adult fish.

2.2. Cas9 MRNA and Guide RNA Synthesis

Synthesis of Cas9 mRNA and guide RNA (gRNA) was performed using previously described methods (Jao et al. 2013; Vejnar et al. 2016). Target gRNA sites were identified using CRISPRscan (Moreno‐Mateos et al. 2015), synthesized using the T7 MEGAscript kit (Invitrogen), and purified by phenol–chloroform extraction and ethanol precipitation. A zebrafish codon‐optimized Cas9 (pT3TS‐nCas9n, Addgene plasmid # 46757) was linearized with XbaI overnight, and 1 µg of linear template DNA was used for in vitro transcription of mRNA using the T3 mMESSAGE mMACHINE (Invitrogen) kit and purified by phenol–chloroform extraction and ethanol precipitation. The following gRNA target sites were used, with the protospacer‐adjacent motif (PAM) site underlined:

  • 5′ GAAGAAGCTGGAGGCGGCTGAGG 3′ for stmn2a, located in exon 3.

  • 5′ GAGGCACGCAAGAACATCATGGG 3′ for stmn2b, located in exon 2.

2.3. CRISPR/Cas9 Mutagenesis and Screening for Founder Lines

Stmn2a , stmn2b, and double stmn2a/b KOs were generated using methods previously described by our laboratory (Armstrong et al. 2016). Cas9 mRNA (100 ng/µL) and gRNA (100 ng/µL) were co‐injected in ∼2 nL volumes into zebrafish embryos at the one‐to‐two‐cell stage of development. DNA cutting efficiency was evaluated through high resolution melting (HRM) analysis of 24 one‐to‐two‐day postfertilization (1–2 dpf) embryos per gRNA target (Erali and Wittwer 2010). The most efficient gRNA target sites were selected for the creation of individual zebrafish lines with disruption of stmn2a and stmn2b reading frames and raised as F0s. Adult F0s were outcrossed with wild type zebrafish and fertilized embryos were screened for indel transmission using HRM.

HRM primer sequences for verification of stmn2a line:

  • Forward: 5′ GGCCCACAGCATCACCTCT 3′

  • Reverse: 5′ CATTTGTCTGTGTTTGCACTAACCC 3′

  • Primer sequences for verification of stmn2b line:

  • Forward: 5′ CGCAGCATACAAAGAGAAGATGAAGG 3′

  • Reverse: 5′ TCCTTCTCCACTGTGTCTCATTACTGG 3′

2.4. General Screening and KO Line Selection

Adult F1s zebrafish were individually separated and anesthetized with 0.02% tricaine (MS‐222, Sigma) and a caudal fin clip was used to extract DNA. DNA was extracted (Extract‐N‐Amp Tissue PCR Kit, Millipore Sigma), and target sites were amplified by polymerase chain reaction (PCR) and followed by restriction fragment‐length polymorphism analysis (BbvCI and NlaIII for stmn2a and stmn2b, respectively). Sequencing (Genome Québec) was performed on amplicons to identify the specific mutation of each founder line. Founder zebrafish carrying frameshift mutations resulting in premature stop codons were selected, and homozygous carriers of each mutation were generated by incrossing F2s to generate our stmn2a KO, stmn2b KO, and double stmn2a/b KO lines. Transgenic Tg[Hb9:GFP] fish, expressing GFP in their motor neurons, were crossed into the KO models to generate a double stmn2a−/− ; stmn2b−/− ; Tg[Hb9:GFP] line.

Primer sequences for verification of stmn2a line:

  • Forward: 5′ TAAAAGGGCCTCAGGTCAGG 3′

  • Reverse: 5′ GTTCTCAATGTGGCTGTCTTTTCC 3′

  • Primer sequences for verification of stmn2b line:

  • Forward: 5′ GTACCCTCTGGCTTAAAACGTAGATG 3′

  • Reverse: 5′ CCTCTCACTCATGAAACGAGCTTG 3′

2.5. In Vitro Feralization (IVF)

The procedure for IVF was adapted from methods previously developed for zebrafish (Draper and Moens 2009). Briefly, at 9 a.m., a double stmn2a−/−; stmn2b−/− adult female zebrafish was anesthetized using 0.02% MS‐222 (Sigma‐Aldrich) in system water. Female zebrafish were blotted dry and placed upside down into the crease of a small sheet (4 cm × 6 cm) of folded laminated plastic paper. Eggs were squeezed out by applying light pressure with the thumb dorsal to the cloaca. Ejected eggs were removed, using a spatula, to a 35 mm petri dish. Eggs from three females were pooled to increase the number of eggs and chance of obtaining fertilized embryos. Male double stmn2a−/−; stmn2b−/− adult (either containing the Hb9:GFP transgene or without it) zebrafish were similarly anesthetized and placed upside down into a foam holder. Sperm was extracted under the microscope by squeezing from the area anterior to the cloaca to the cloaca itself using a pair of fine‐tipped untextured forceps (Fisher Scientific). Good quality sperm (white and not watery) was collected using an aspirator equipped with a gel loading tip (0.6 mm O.D., Fisher Scientific) marked at every µL. Three or more males were pooled to obtain a minimum of 1 µL of sperm. Eggs were fertilized by adding the sperm and 300 µL of system water for 5 min at room temperature (RT) (∼23°C). Then 4 mL of system water was added, and eggs were incubated for 3 h at 28.5°C. After this, fertilized eggs were transferred to 40 mL culture bottles and raised at 28.5°C. IVF‐derived embryos were used for the survival assay, NMJ electrophysiological recordings, and motor axon transection and repair assays.

2.6. Motor Function

Motor function was assessed using methods previously described (Armstrong and Drapeau 2013a). Individual zebrafish aged 52–54 hours postfertilization (hpf) were placed in the center of an aquatic arena (150 mm petri dish) containing fresh system water that was temperature controlled between 24°C and 25°C. Individual zebrafish were allowed to habituate to the environment for 30–60 s. A light touch to the tail was applied using a pair of forceps, evoking a burst swimming motor escape response. Movement was recorded from above at 30 Hz (Grasshopper 2 camera, Point Gray Research) until individual fish either stopped moving or until they reached the side of the arena. Total swim distance and mean swim velocity were quantified using manual movement tracking (Manual Tracking plugin, ImageJ).

2.7. RNA Extraction and Quantitative Real‐Time PCR (qRT‐PCR)

RNA was extracted from 30 pooled 2 dpf embryos using the trizol‐chloroform method. Purity and quality of RNA were verified using a NanoDrop instrument. 2.5 µg of RNA was used to create a cDNA library using the SuperScript VILO cDNA Synthesis Kit (Invitrogen). Quantitative real‐time PCR was performed on the cDNA libraries using the SYBR Green Supermix (Bio‐Rad Laboratories). The delta–delta Ct method of relative quantification was performed with gapdh as an internal control (Livak and Schmittgen 2001).

Primer sequences for RT‐qPCR analysis of stmn2a line:

  • Forward: 5′ GAACAAGGAAAACCGTGAGG 3′

  • Reverse: 5′ CTTGTTCCTGCGGACGAT3′

  • Primer sequences for RT‐qPCR analysis of stmn2b line:

  • Forward: 5′ CTGAAAGCCATGGAGGAGAA 3′

  • Reverse: 5′ ACCAGCTGAGCGTGCTTC 3′

  • Primer sequences for gapdh:

  • Forward: 5′ CGCTGGCATCTCCCTCAA 3′

  • Reverse: 5′ TCAGCAACACGATGGCTGTAG 3′

2.8. NMJ Colocalization and Axonal Microtubule Networks

Immunofluorescence microscopy experiments were performed to assess NMJ integrity. Double labeling of presynaptic and postsynaptic membranes was achieved using a synaptotagmin 2 (Syt2, presynaptic) antibody (DSHB, znp‐1‐s, mouse) and sulforhodamine‐conjugated alpha‐bungarotoxin (α‐Btx, postsynaptic) marker (Millipore Sigma, T0195) as previously described (Armstrong and Drapeau 2013a). Briefly, 10 dechorionated 2 dpf zebrafish were fixed in 4% paraformaldehyde (PFA) in phosphate‐buffered saline solution (PBS) overnight at 4°C with a gentle rotation. Fixed zebrafish were then rinsed with PBS (3 × 15 min) and incubated with 1 mg/mL collagenase in PBS solution for 45 min at RT. Zebrafish were again rinsed in PBS (3 × 15 min) and incubated in PBST (PBS, Triton X‐100) for 30 min at RT and then incubated in a 10 mg/mL sulforhodamine‐conjugated α‐Btx in PBST solution for 30 min at RT. Following this, zebrafish were rinsed with PBST (3 × 15 min) and incubated for an hour in fresh blocking solution (2% Goat serum, 1% bovine serum albumin [BSA], 0.1% Triton X‐100, and 1% DMSO in PBS) at RT before being left overnight in block solution containing the Syt2 primary antibody (1:100). The next day, zebrafish were rinsed with PBST (4 × 15 min) and incubated at RT for 5 h in fresh blocking solution containing the secondary antibody (Alexa fluor 488, Abcam, Ab150117, goat anti‐mouse, 1:2000). Zebrafish were rinsed with PBST (3 × 15 min) and incubated in PBST overnight at 4°C with a gentle rotation and then placed in a solution containing 70% glycerol and mounted on glass slides for imaging. NMJs were visualized with a 60×/1.42 oil immersion objective on a Quorum Technologies microscope with an 89 NORTH LDI spinning disk confocal mounted on an Olympus BX61W1 fluorescence microscope, connected to a photometrics prime BSI camera. Images were acquired using Volocity software (Improvision).

2.9. Electrophysiology

Using previously described methods (Buss and Drapeau 2002), zebrafish aged 50–54 hpf were anaesthetized in 0.02% tricaine (Sigma) dissolved in modified Evans solution containing, in mM: 134 NaCl, 2.9 KCl, 2.1 CaCl2, 1.2 MgCl2, 10 HEPES, and 10 glucose, adjusted to 290 mOsm and pH 7.8. Zebrafish were then pinned using fine (0.001 in.) tungsten wires through their notochords to a Sylgard‐lined dish. The outer layer of skin between the pins was removed using a fine glass electrode and forceps, exposing the trunk musculature. The preparation was then visualized by oblique illumination (Zeiss Examiner A1 upright microscope), and standard whole‐cell voltage clamp recordings were obtained from fast‐twitch (embryonic white) muscle cells identified visually by their oblique orientation to the body axis. Glass electrodes of 5–6 MΩ were pulled from thin‐walled Kimax‐51 borosilicate glass (Kimble Chase) and filled with intracellular solution (in mM: 130 CsCl, 2 MgCl2, 10 HEPES, and 10 EGTA adjusted to pH 7.2, 290 mOsm). Cells were held near their resting potential at −65 mV, and series resistance was <8 MΩ and compensated to 70%–90%. Tetrodotoxin (TTX, Tocris Bioscience, 1069) of 1 µM was perfused over the preparation to isolate spontaneous (quantal) miniature endplate currents (mEPCs) over a 10‐min recording period. All electrophysiological data were sampled at 40 kHz using an Axopatch 200B amplifier (Molecular Devices) and digitized using a Digidata 1550B (Molecular Devices) and later analyzed using pCLAMP 10 software (Molecular Devices).

2.10. Axon Transection and Recovery

Wild type and double stmn2a −/−; stmn2b −/− zebrafish expressing a single copy of the Tg[Hb9:GFP] transgene were selected for axon ablation. Individual zebrafish were embedded in 1% low melting point agarose, prepared using system water containing 0.02% tricaine (Sigma), at 50 hpf. Pre‐transection images of fluorescently labeled ventral root axon projections were collected. A microinjection needle was then used to fully transect an individual ventral root axonal projection in a single hemisomite between somites 13 and 17. Cut ventral roots were reimaged every hour for 24 h using a 60×/1.42 water immersion objective.

2.11. Statistical Analyses

Statistical analyses were performed using Prism 9 (GraphPad Software Inc.). Shapiro–Wilk test was used to test for normal sample distributions. Tests with only two groups were compared using unpaired students’ t‐tests or Mann–Whitney rank sum tests when data were not found to be normally distributed. Kruskal–Wallis tests were used to compare data sets containing more than two non‐normally distributed samples, followed by Dunn's post hoc multiple comparisons test. Chi‐squared tests were used to compare genotypic frequencies. Data are reported as mean ± standard error measure (e.g., black lines on figures) or specific numerical values in the text. Significance was assessed at p < 0.05.

3. Results

3.1. Generation of stmn2a, stmn2b, and Double KO Zebrafish Lines

The gene encoding STMN2 is represented by two orthologs in the zebrafish genome: stmn2a and stmn2b (Figure 1A). Guide RNAs targeting the coding regions within exons 2 and 3 for each gene were individually evaluated for cutting efficiency using the CRISPR/Cas9 mutagenic system. HRM analysis of F1 offspring revealed that a gRNA targeting exon 3 of stmn2a and a gRNA targeting exon 2 of stmn2b were robustly cutting DNA at these loci and were subsequently selected for generating our mutant fish lines. Sanger sequencing of F2 homozygous fish revealed an 11‐nucleotide deletion in exon 3 of stmn2a and a 13‐nucleotide deletion in exon 2 of stmn2b (Figure 1B), resulting in premature stop codons in the open reading frame of their respective transcripts that could be screened by the loss of restriction enzyme sites in amplified PCR products (Figure 1C). A double stmn2a and stmn2b KO fish line was generated by incrossing both lines. Additionally, a double KO fish line expressing the Tg[Hb9:GFP] transgene (Flanagan‐Steet et al. 2005) was also crossed into our mutant lines to create a double KO and Tg[Hb9:GFP] transgenic fish line. To confirm that the mutations were disrupting gene function, we quantified stmn2a and stmn2b transcript levels using cDNA synthesized from RNA collected from pooled 2 dpf zebrafish. Both homozygous mutant lines showed significantly reduced expression levels of stmn2a and stmn2b transcripts, suggesting that the transcripts were removed through nonsense‐mediated decay, indicating that both mutations likely confer a loss‐of‐function phenotype (Figure 1D). HRM analysis of potential CRISPR off‐target sites did not find amplicons containing indels in double KO fish (Figures S1–S5).

FIGURE 1.

FIGURE 1

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Development of the stmn2a and stmn2b KO zebrafish lines. (A) Schematic representation of zebrafish stmn2a and stmn2b genes showing gene structure. (B) Location of guide RNAs (pink lines) for each gene and mutant lines selected for use. An 11‐nucleotide deletion and a 13‐nucleotide deletion were produced in stmn2a and stmn2b, respectively, resulting in premature stop codons in the open reading frame. (C) Loss of BbvCI (stmn2a) and NlaIII (stmn2b) restriction enzyme digestion sites at their respective gRNA target sites was used for genotyping of digested amplicons. (D) Analysis of gene expression using RT‐qPCR of cDNA derived from RNA collected from 2 dpf zebrafish. Expression levels are relative to gapdh. A significant reduction in stmn2a expression in stmn2a−/− zebrafish (p < 0.05) and a significant reduction in stmn2b in stmn2b−/− fish (p < 0.05) were observed, suggesting the mutant transcripts are targeted for nonsense‐mediated decay. No stmn2a compensation was observed in the stmn2b−/− fish, whereas a trend (p = 0.11) towards stmn2b compensation in stmn2a−/− fish was observed. Statistical analysis was performed by Mann–Whitney tests, and significance determined at p < 0.05. Sample sizes: wild type, N = 4 batches of 30 pooled embryos; stmn2a−/−, N = 3; stmn2b−/−, N = 3. (E) A significant reduction in the expected frequency of double stmn2a −/−; stmn2b −/− 2 dpf embryos was identified following a stmn2a +/−; stmn2b ± × stmn2a +/−; stmn2b ± incross when compared to the expected frequency (chi‐squared test, p = 0.003). (F) Survival curve showing a reduction in stmn2a −/−; stmn2b −/− zebrafish survival between 5 and 30 days of development in comparison to wild type (chi‐square test, p = 0.0011), n = 50 larvae for each group *p < 0.05, **p < 0.01.

As both fish Stmn2a and Stmn2b are similar to one another (Figure 2), we speculated that each paralog might be able to compensate for the loss of expression of the other by displaying elevated transcriptional expression. However, examination of transcript levels from whole 2 dpf zebrafish revealed that there was no change in the expression of stmn2a in stmn2b −/− zebrafish and a trend of increased expression of stmn2b in stmn2a−/− cDNA samples, but this did not reach significance (Figure 1D). Examination of genotype frequency following a double heterozygous incross (e.g., stmn2a +/−; stmn2b +/−) revealed that fewer double KO zebrafish (e.g., stmn2a −/−; stmn2b −/−) were surviving to this stage of development (Figure 1E). We next investigated if there were changes in survival over the first 30 days of development. We observed that double KO zebrafish displayed impaired survival when compared to wild type during the first month of development (Figure 1F). Surviving double KO zebrafish were able to reach sexual maturity normally; however, adult male double stmn2a −/−; stmn2b −/− zebrafish were behaviorally unable to fertilize embryos of any genotype (likely due to poor adult motor function) but could be used for IVF to generate clutches of double stmn2a −/−; stmn2b −/− embryos.

FIGURE 2.

FIGURE 2

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Amino acid sequence alignment between human STMN2 and zebrafish Stmn2a (top), human STMN2 and zebrafish Stmn2b (middle), and zebrafish Stmn2a and Stmn2b (bottom). Zebrafish Stmn2a shares 78% identity and 90% similarity with human STMN2, whereas Stmn2b shares 73% identity and 88% similarity with human STMN2. Zebrafish Stmn2a and Stmn2b are 85% identical and 93% similar to each other.

3.2. Stmn2 KO Zebrafish Display a Significant Reduction in Motor Function

Given that both stmn2a and stmn2b transcripts are expressed during development, we next sought to determine whether the loss of expression of either transcript affected motor function. At 2 dpf, zebrafish display burst swimming behavior in response to a light tail touch (Buss and Drapeau 2001). Using zebrafish aged 52–54 hpf, we conducted touch‐response assays in an aquatic arena (Figure 3A,B), following previously established methods (Armstrong and Drapeau 2013a, 2013b). Zebrafish in all genetic groups were able to respond to the light touch to the tail. Examination of motor parameters revealed a significant reduction in mean swim velocity in stmn2a −/−, stmn2b −/−, and double stmn2a −/−; stmn2b −/− fish when compared to controls (Figure 3C). Furthermore, total swim distance was significantly reduced in stmn2a −/− and double stmn2a −/−; stmn2b −/− fish when compared to wild type zebrafish (Figure 3D), whereas stmn2b −/− fish exhibited swim distances that were similar to wild type controls (Figure 3D).

FIGURE 3.

FIGURE 3

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Two‐day old zebrafish touch‐evoked motor response assays for stmn2a −/−, stmn2b −/−, and double stmn2a −/−; stmn2b −/− fish. (A) Representative images of 2 dpf zebrafish. Scale bar represents 400 µm. (B) Five individual superimposed touch‐evoked motor traces from wild type, stmn2a −/−, stmn2b −/−, and double stmn2a −/−; stmn2b −/− genetic groups. (C) Mean swim velocity ± standard error and sample sizes were as follows: wild type (0.55 ± 0.03 cm/s, n = 30); stmn2a −/− (0.9 ± 0.02 cm/s, n = 30); stmn2b −/− (0.39 ± 0.02 cm/s, n = 45); and double stmn2a −/− and stmn2b −/− (0.22 ± 0.01 cm/s, n = 39). Statistical differences were as follows: wild type versus stmn2a −/−, p < 0.0001; wild type versus stmn2b −/−, p = 0.004; wild type versus double stmn2a −/− and stmn2b −/−, p < 0.0001. (D) Mean swim distance ± standard error and sample sizes were as follows: wild type (12.4 ± 1.4 cm, n = 30); stmn2a −/− (5.7 ± 0.8 cm, n = 30); stmn2b −/− (9.3 ± 0.8 cm, n = 45); double stmn2a −/− and stmn2b −/− (6.3 ± 0.7 cm, n = 39). Statistical differences were as follows: wild type versus stmn2a −/−, p = 0.0006; wild type versus double stmn2a −/− and stmn2b −/−, p = 0.0009. Data points in (C) and (D) represent individual zebrafish measurements with black/grey bars representing the overall group mean ± standard error. Kruskal–Wallis multiple comparisons tests were used to determine significance. *p < 0.05, **p < 0.01, and ***p < 0.001 and ****p < 0.0001.

3.3. Double stmn2a −/−; stmn2b −/− Zebrafish Present Decreased Colocalization of Pre‐ and Postsynaptic NMJ Markers

As motor function impairment could result from abnormal NMJ structure, we next investigated if there were defects at these sites. Emanating from the spinal cord, ventral roots in 2‐day‐old zebrafish are composed of 20–30 axons from primary and secondary spinal motor neurons (Myers 1985; Westerfield et al. 1986) that project to and innervate axial muscles in each hemisegment. Reduced NMJ integrity has been described in previous ALS‐mutant zebrafish models, where receptors can be considered “orphaned” when postsynaptic puncta (e.g., nicotinic acetylcholine receptors, AchR) do not colocalize with presynaptic motor neuron markers (e.g., synaptotagmin 2, Syt2) (Armstrong and Drapeau 2013b; Bose et al. 2019). To investigate trunk NMJs in our mutant stmn2 models, we performed immunofluorescence confocal imaging of fixed zebrafish (aged 55–56 hpf) and labeled pre‐ and postsynaptic NMJ components with an antibody targeting Syt2 and alpha‐bungarotoxin conjugated to sulforhodamine (aBtx) targeting AchRs (Figure 4A). We observed a significant increase in the number of orphaned presynaptic Syt2 puncta in stmn2a −/− and double stmn2a −/−; stmn2b −/− zebrafish, suggesting that the motor axons in these zebrafish possessed axon terminals that did not project to AchRs clusters (Figure 4B). Conversely, we observed the number of orphaned postsynaptic AchR puncta to be slightly increased in our double stmn2a −/−; stmn2b −/− fish model when compared to wild type NMJs (Figure 4C). To further investigate the nature of NMJ function, we recorded mEPCs on fast‐twitch muscle cells of the zebrafish trunk (somites 10–15). In these recordings, we observed that mEPC amplitude in double stmn2a −/−; stmn2b −/− zebrafish was reduced in comparison to recordings from aged‐matched wild type fish (Figure 4D,E), with no change being observed in the frequency of mEPCs (Figure 4F). Additionally, we observed no differences between wild type and double stmn2a −/−; stmn2b −/− zebrafish in our measures of mean 10%–90% rise time (1.35 ± 0.19 vs. 1.75 ± 0.09 ms, p = 0.47), mean decay (τ) constant (13.52 ± 1.44 vs. 12.99 ± 2.30, p = 0.20), mean resting muscle membrane potential (−62.75 ± 2.71 vs. −65.83 ± 2.51 Vm, p = 0.42), or mean muscle membrane capacitance (69.38 ± 12.34 vs. 76.02 ± 8.05 pF, p = 0.35).

FIGURE 4.

FIGURE 4

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Two‐day‐old zebrafish NMJs from double stmn2−/− and stmn2b−/− zebrafish display increased pre‐ and postsynaptic puncta and decreased quantal size of mEPCs. (A) Representative immunofluorescent images of zebrafish NMJs from wild type and double stmn2a−/− and stmn2b−/− embryos. Scale bars represent 25 µm. Quantification of orphaned presynaptic Syt2 puncta (red arrowheads) (B) and postsynaptic AchR puncta (green arrows) (C). Mean number of orphaned presynaptic terminals ± standard error and sample sizes were as follows: wild type (7.7 ± 0.9 terminals, n = 13); stmn2a−/− (10.7 ± 0.7 terminals, n = 18); stmn2b−/− (8.4 ± 1.1 terminals, n = 14); double stmn2a−/− and stmn2b−/− (11.7 ± 0.6 terminals, n = 16). Mean number of orphaned postsynaptic terminals ± standard error and sample sizes were as follows: wild type (4.2 ± 0.9 terminals, n = 13); stmn2a−/− (5.1 ± 0.5 terminals, n = 18); stmn2b−/− (4.9 ± 0.7 terminals, n = 14); double stmn2a−/− and stmn2b−/− (6.4 ± 0.5 terminals, n = 16). Statistical differences in (B) were as follows: wild type versus stmn2a −/−, p = 0.047; wild type versus double stmn2a −/− and stmn2b −/−, p = 0.004. Statistical difference in (C) was as follows: wild type versus double stmn2a −/− and stmn2b −/−, p = 0.011. (D) Example mEPC events in wild type and double stmn2a−/− and stmn2b−/− fish. (E) Quantification of mean mEPC amplitude. Mean mEPC amplitude was found to be reduced (p = 0.003) in double stmn2a−/− and stmn2b−/− zebrafish in comparison to recordings obtained from wild type fish. (F) Quantification of mean mEPC frequency. No significant difference was observed between wild type and double stmn2a−/− and stmn2b−/− groups (p = 0.148). Samples sizes for mEPC recordings were as follows: wild type = 13; double stmn2a−/− and stmn2b−/−  = 10. Data points represent individual means from zebrafish recordings with black bar representing the overall group mean ± standard error. Kruskal–Wallis multiple comparisons tests were used to determine significance (B and C) or a t‐test (E and F). mEPCs, miniature endplate currents. *p < 0.05, **p < 0.01.

3.4. Double stmn2a −/−; stmn2b −/− Zebrafish Display Impaired Nerve Regeneration Following Ventral Root Nerve Injury

As ventral root projections in our double stmn2a and stmn2b KO zebrafish only displayed minor NMJ defects, we next explored if there were abnormalities in axon regeneration following injury. Using restrained zebrafish at age 50 hpf, we transected single ventral root projections using a fine glass microelectrode and then recorded axon regeneration over a 24‐h period. Ventral root axon projections were visualized using fish expressing the Tg[Hb9:GFP] transgene (Figure 5A). Wild type zebrafish expressing the Tg[Hb9:GFP] transgene displayed fully degenerated distal sections of the ventral root projection within 3–4 hours post injury (hpi) (Figure 5A). However, in double stmn2a−/−; stmn2b−/− zebrafish expressing the Tg[Hb9:GFP] transgene (Figure 5B), transected ventral root project took longer to fully degenerate (6–12 hpi). Examination of the ventral root projection at 24 hpi revealed that 7/8 wild type motor neurons had fully regrown their axon projections, whereas none of our double stmn2a−/−; stmn2b−/− zebrafish were able to regrow their ventral root axon projections (Figure 5C). These findings suggest a significant role for Stmn2a/b in motor axon maintenance and regeneration in zebrafish embryos.

FIGURE 5.

FIGURE 5

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Delayed ventral root motor axon degeneration following axotomy in 2 dpf stmn2a −/−; stmn2b −/− zebrafish. (A) Example confocal images of ventral root motor axons of a wild type (top) expressing Tg[Hb9:GFP] and stmn2a−/− ; stmn2b−/− zebrafish (bottom) expressing Tg[Hb9:GFP] over a 24 h period following injury (hpi). White arrows indicate location of axotomy. Scale bar represents 25 µm. (B) Quantification of the length of time it took for distal motor axon degeneration to occur. Axon degeneration time was significantly longer in stmn2a−/− ; stmn2b−/− fish when compared to wild type zebrafish. Sample sizes are as follows: wild type, n = 8; stmn2a−/− ; and stmn2b−/− , n = 7. Data points represent individual axon degeneration times from individual zebrafish with black bar representing the overall group mean ± standard error. A t‐test was used to make statistical comparisons. ***p < 0.001. (C) Tabulation of the number of ventral roots regrown at 24 hpi.

4. Discussion

Using CRISPR/Cas9 mutagenesis, we developed a zebrafish stmn2 KO model and investigated abnormalities in the motor system during zebrafish development. To generate our KO model, we selected mutant zebrafish lines containing an 11‐nucleotide deletion in exon 3 of stmn2a and a 13‐nucleotide deletion in exon 2 of stmn2b, both of which produce premature stop codons that resulted in their mRNA transcripts being targeted by nonsense‐mediated decay (Figure 1D). Single‐cell RNA‐Seq data (Sur et al. 2023) and in situ hybridization experiments (Burzynski et al. 2009) suggest that both stmn2a and stmn2b are expressed in the developing zebrafish embryo. To examine if either stmn2a or stmn2b could compensate for loss of the other, we examined levels of their transcripts in our single gene KO lines. We found no changes in the levels of stmn2a in our stmn2b −/− line; however, there was a trend of increased stmn2b transcript levels in our stmn2a −/− line. Although either stmn2a or stmn2b single gene KO models did not show reduced survival during development, double stmn2a−/−; stmn2b−/− zebrafish did display reduced survival (Figure 1E,F). Despite this early survival defect in our double stmn2a−/−; stmn2b−/− model, we were able to raise some fish to adulthood. However, we observed that males were behaviorally unable to fertilize female zebrafish of any genotype. IVF using double stmn2a−/−; stmn2b−/− sperm confirmed the male gametes were able to fertilize eggs; however, their consistent inability to breed naturally is noteworthy and likely reflects impaired motor function. A limitation of our study was that we were unable to attempt rescue experiments (i.e., blastocyst injection of wild type stmn2a or stmn2b mRNA) in double stmn2a −/− and stmn2b −/ − zebrafish due to the fragility of IVF‐derived embryos.

As both orthologs of stmn2 are expressed during development and are thought to play a role in microtubule stability in motor axons, we next investigated if there were abnormalities in larval motor function (Figure 3). Although all our models were able to respond to a light touch to the tail evoking a motor touch response, we did observe that stmn2a−/− , stmn2b−/− , and double stmn2a−/− ; stmn2b−/− zebrafish displayed a reduction in mean swim velocity, whereas only our stmn2a−/− and double stmn2a−/−; stmn2b−/− models displayed a reduction in total swim distance. These findings suggest that while not essential for motor function, loss of Stmn2 expression is coincident with a slight impairment in motor function at embryonic stages of zebrafish development. To investigate potential causes of impaired motor function, we next investigated if there were abnormalities at NMJs. Examination of presynaptic and postsynaptic components of the NMJ revealed that double stmn2a−/− ; stmn2b−/− zebrafish possessed more pre‐ and postsynaptic puncta labeling (Figure 4). The number of pre‐ and postsynaptic NMJs that did not colocalize with each other, however, was increased in our double stmn2a−/− ; stmn2b−/− model at 2 dpf. Our single gene KO models displayed NMJs that were similar to wild type with one exception being that stmn2a−/− trunks displayed an increase in the number of orphaned presynaptic puncta. To further investigate NMJ function, we recorded mEPCs from fast‐twitch muscle cells. We observed a reduction in the amplitude of mEPCs in double stmn2a−/− ; stmn2b−/− zebrafish and no change in any other metric of mEPCs when compared to wild type recordings (Figure 4D–F). This suggests that although NMJs are able to be formed in our model, the synaptic strength of individual NMJs is reduced. An alternative possibility is that there is a defect in the clustering of AchRs on muscle cells in our double stmn2a−/−; stmn2b−/− model. Muscle cell abnormalities, including the presence of increased centrally located nuclei (suggesting muscle cell regeneration) and fragmented NMJs have been reported in Stmn2 KO mice (Guerra San Juan et al. 2022). As both stmn2a and stmn2b are widely expressed in the embryo and larval CNS, it maybe that other cell types, in particular, neurons that process sensory information, are impaired. However, none of our zebrafish failed to react to the tail touch, suggesting that at this stage in development the sensory system is intact.

In a final series of experiments, we investigated if there were defects in motor axon regeneration following injury in our double stmn2a−/−; stmn2b−/− model. Following motor axon transection, we observed a slower time course for distal regions of the motor axon to degenerate (Figure 5). In addition, we observed an impaired ability to regrow motor axons over a 24‐h period in our double stmn2a−/−; stmn2b−/− model when compared to wild type motor axons that displayed a near complete ability to regrow motor axons. Recently, it was shown that Stmn2 is essential for motor axons to reinnervate NMJs following sciatic nerve crush in mice (Beccari et al. 2025). The findings made in our study support the notion that Stmn2 is involved with axon regrowth following injury.

Recent advances in our understanding of misregulated genes in the neurodegenerative disorder ALS have brought increased attention to the role of STMN2 in disease. In motor neurons from individuals with ALS, the RNA‐binding protein TDP‐43 is mislocalized from the nucleus to the cytoplasm where it can form insoluble aggregates (Neumann et al. 2006). TDP‐43 (encoded by TARDBP) is a ubiquitously expressed member of the heterogeneous nuclear ribonucleoprotein family of proteins that is involved in several steps of RNA processing, including of the STMN2 transcript. In healthy motor neurons, the full‐length STMN2 transcript is composed of 5 exons, but following depletion of TDP‐43 or expression of ALS‐associated variants of TDP‐43, a cryptic exon in the first intron of STMN2 termed exon 2a is incorporated in the pre‐mRNA that includes a premature polyadenylation site as well as TDP‐43 binding motifs (Klim et al. 2019; Melamed et al. 2019; Baughn et al. 2023). This aberrant splicing of the STMN2 transcript results in the expression of a severely truncated and nonfunctional protein and reduces expression of full‐length STMN2 (Baughn et al. 2023). Although zebrafish and murine genomes do not possess the corresponding cryptic STMN2 exon 2a, these comparative models can serve as useful tools for investigating the consequences that loss of stmn2 expression has on nervous system function in vivo. In summary, our findings demonstrate that loss of stmn2a and stmn2b expression results in a moderate motor phenotype in zebrafish and impairment in the ability to regenerate motor axons following injury.

Authors Contributions

Experimental data collection and analysis were performed by Tyler J. N. Gurberg, Ziyaan A. Harji, Christian J. Rampal, Joséphine Sacy‐Richer, Amy Wang, Esteban C. Rodríguez, and Gary A. B. Armstrong. Tyler J. N. Gurberg and Gary A. B. Armstrong wrote the original draft manuscript. Draft revisions were completed by Gary A. B. Armstrong. Funding for the project was obtained by Gary A. B. Armstrong.

Funding

This research was supported by a Natural Sciences and Engineering Research Council of Canada Discovery Grant (G.A.), a Canadian Institutes of Health Research CGS M and Project Grant (T.G. and G.A.), and an ALS Canada—Brain Canada Discovery Grant (G.A.).

Disclosure

All data pertain to the creation of this manuscript are contained within it.

Ethics Statement

All experiments were approved by the Montreal Neurological Institute and Hospital's Animal Care Committee (ACC) and followed the guidelines set by the Canadian Council for Animal Care (CCAC). Animal care protocol approval # MNI‐7890.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supplementary Material: dneu70054‐sup‐0001‐Figures.docx

DNEU-86-0-s001.docx (2.3MB, docx)

Gurberg, T. J. N. , Harji Z. A., Rampal C. J., et al. 2026. “Impaired Motor Function in a Zebrafish Stathmin‐2 Knockout Model.” Developmental Neurobiology 86, no. 4: e70054. 10.1002/dneu.70054

A manuscript preprint has been uploaded to bioRxiv: (Gurberg et al., 2025)

This article is based upon M.Sc. thesis research conducted by the first author: Tyler Gurberg.

Data Availability Statement

Data supporting these findings are available from the corresponding author by request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Material: dneu70054‐sup‐0001‐Figures.docx

DNEU-86-0-s001.docx (2.3MB, docx)

Data Availability Statement

Data supporting these findings are available from the corresponding author by request.

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