Ependymoglial cells are critical for cortex regeneration in axolotls

Domain: neuroscience
Published: 2026-01-20
Editorial outcome: major_revision

Abstract
Developing precise targeted cell ablation in the axolotl (Ambystoma mexicanum) is crucial for elucidating the roles or interactions of specific cell types in regeneration and modeling diseases. Here we establish a Nitroreductase (NTR)-based inducible cell ablation system in axolotls. Through generation of Sox2:Cherry-NTR knock-in axolotls, we achieve efficient ablation of ependymoglial cells (EGCs) in the central nervous system. Combined spinal cord and brain transplantation and injury models demonstrate regeneration failure upon EGC depletion, suggesting that EGCs are sole source of central nervous system regeneration. Additionally, EGC ablation in the spinal cord resulted in delayed tail regeneration. Moreover, we establish NeuroD6:Cherry-NTR and NeuroD6:Cherry-NTR 2.0 knock-in lines to ablate postmitotic cortical neurons, enable the investigation of brain regeneration after large-scale neuronal depletion. We found that NTR 2.0 (but not NTR) leads to elimination of >95% of targeted neurons in the dorsal pallium. All lost neuronal subtypes are chronologically regenerated with laminar-like distribution mirroring developmental patterning. Finally, we create Cre-LoxP-based conditional NTR 2.0 transgenic axolotls using a constitutive CAGGs promoter, enabling tissue-specific ablation of specific cell types when crossed to existing Cre lines. In summary, our study establishes an efficient and versatile targeted cell ablation system in axolotls, providing a valuable tool for deep dissection of tissue regeneration in axolotls.

Introduction
Tissue and organ regeneration represent a central challenge in the fields of biology and medicine. It requires precise coordination of diverse cell populations to restore structural integrity and functional capacity. Although significant advances in understanding the mechanisms underlying tissue regeneration have been achieved through studies across various species in recent years1–8, many key issues, such as the precise roles of each cell type and their potential interactions during complex tissue regeneration, remain largely elusive. Such knowledge gaps hinder the development of targeted therapeutic strategies to improve mammalian tissue repair in regenerative medicine.

The axolotl (Ambystoma mexicanum) has emerged as a premier model for regeneration research due to its unparalleled ability to regenerate organs such as limbs9–17, spinal cord18–24, and brain tissue25–31. Successful regeneration of injured organs relies on the interplay of various cell types, each contributing in either a direct, indirect, or dual manner1,32–37. Cells with direct roles participate in regeneration via proliferation and differentiation to construct lost tissues. Cell fate mapping using genetic approaches or tissue transplantation has revealed that lineage-restricted progenitors are the primary contributors to limb regeneration11,38–40. Conversely, certain cells indirectly influence regeneration by modulating the behavior of neighboring cells. It has been reported that macrophages interact with limb blastema cells to regulate their proliferation and differentiation by shaping the inflammatory microenvironment41–46. Additionally, nerves and the apical epithelial cap release signaling molecules critical for guiding limb regeneration13,14,32,47–53. Surgical removal of nerves or the apical epithelial cap, or chemical depletion of macrophages, blocks limb regeneration42,48,54–56, whereas muscle tissue is dispensable57.

Recent advances in multi-omics and single-cell sequencing have identified versatile cell types that interact dynamically during regeneration. In Xenopus tail regeneration, “regeneration-organizing cells” were discovered to coordinate the regenerative process58. However, precise ablation of specific cell types for investigation of their roles remains challenging due to limitations in surgical or chemical methods, including imprecision, inaccessibility, and off-target effects.

Genetic conditional cell ablation systems, such as diphtheria toxin receptor/toxin59,60, herpes simplex virus thymidine kinase/ganciclovir61,62, nitroreductase/metronidazole (NTR/MTZ)63–66, and Kid/Kis67, have been widely applied in model organisms. The NTR/MTZ system, originally isolated from Escherichia coli, catalyzes the conversion of non-toxic MTZ into cytotoxic DNA cross-linking agents, inducing apoptosis in NTR-expressing cells64,66. This system has been extensively utilized in zebrafish63–66,68,69. By driving NTR expression under tissue-specific promoters in transgenic zebrafish, targeted cell ablation is achieved upon MTZ exposure. The system’s reversibility (“on/off” capability) further enhances its utility63–66. Combined with tissue-specific promoters or genetic knock-in approaches, the NTR/MTZ system provides an easy, efficient, and highly specific ablation method.

The brain, as the most complex organ, poses unique challenges for repair following injury or degenerative disorders. Axolotls exhibit remarkable brain regeneration after trauma26–28, making them an ideal model for brain regeneration research. While ependymoglial cells (EGCs) have been established as crucial players in central nervous system (CNS) regeneration24,27,28,70–73, significant gaps remain in understanding their precise contributions involved in this process. Moreover, existing studies predominantly focus on physical injury or drug-based models, which lack the precision for targeting specific neuronal subtypes. The Simon lab previously induced a Parkinson’s-like phenotype in newts by ablating midbrain dopamine neurons with 6-hydroxydopamine31,71. Thus, developing genetic-based neuronal ablation methods in regenerative contexts is imperative.

Here, we establish an NTR-based ablation system in axolotls. Using the endogenous Sox2 or NeuroD6 promoter to drive NTR expression, we achieve efficient ablation of EGCs and cortical neurons. We demonstrate that the regeneration of the spinal cord and brain is completely abolished upon EGC depletion. Following large-scale neuronal ablation, we found that neuronal regeneration is organized in a chronological, laminar-like distribution that recapitulates patterns of developmental neurogenesis. Furthermore, we demonstrate the superiority of NTR 2.0 over NTR and develop Cre-loxP-based reporter NTR 2.0 lines compatible with any Cre line for targeted ablation. This toolkit advances the dissection of cellular interactions in regeneration, addressing critical gaps in axolotl regenerative biology.

Results
Effective ablation of Sox2+ cells in the lateral line, but not in the CNS, with MTZ in Sox2:Cherry-NTR knock-in axolotls
To develop targeted cell ablation in axolotls using the NTR system, we targeted EGCs, which have previously been established as the potential cellular source of spinal cord19,23,24 and brain regeneration26–28,72 in salamanders. However, their precise contribution to CNS regeneration remains to be fully elucidated. To address this gap, we attempted EGC ablation in axolotls and investigated their contributions to CNS regeneration and interactions with surrounding tissues during complex tissue regeneration. SOX2 is a well-recognized marker of EGCs24,74, we therefore created a Sox2:Cherry-NTR knock-in line using a semi-homology-directed recombination method by inserting the Cherry-NTR coding sequence into the axolotl Sox2 genomic locus (Supplementary Fig. 1a). Consistent with the previous observations38,75, CHERRY expression was detected in the noses, eyes, spinal cord, brain, and lateral line of Sox2:Cherry-NTR larvae (Supplementary Fig. 1b, c). Immunofluorescence confirmed co-localization of CHERRY and SOX2 in lateral line neuromasts (Supplementary Fig. 1d, e), spinal cord (Supplementary Fig. 1d, f), and brain (Supplementary Fig. 1g, h), indicating faithful expression of CHERRY-NTR in Sox2+ cells.

We then assessed ablation of SOX2+ cells in Sox2:Cherry-NTR axolotls. We observed that the intact neuromast is a cup-shaped structure predominantly composed of SOX2+ cells, surrounded by a ring of weakly SOX2+ cells (Supplementary Fig. 2a and Supplementary Movie 1). Following MTZ treatment, the CHERRY fluorescent signals in the neuromasts of Sox2:Cherry-NTR became scattered at 2 and 5 days post treatment (dpt), compared to controls exposed to DMSO (Fig. 1a, b). Immunofluorescence staining of sections at 3 dpt showed that the neuromasts of MTZ-treated animals have a disrupted structure and a significant reduction of both the CHERRY and SOX2 signals (Fig. 1c, d), compared to the controls. Time-lapse live imaging, along with section and whole-mount analyses, showed that CHERRY+ SOX2+ cell ablation and neuromast destruction progressed over time following MTZ treatment (Supplementary Fig. 2b–e), while some weakly SOX2+ cells adjacent to the cup-shaped structure appeared refractory to ablation (Fig. 1c). Moreover, compared to the controls, apoptotic cells were detected by cleaved Caspase-3 at the neuromast location at 1 dpt (Fig. 1e, f). This was followed by recruitment of macrophages, as indicated by IBA1 immunostaining at 3 dpt (Fig. 1g, h). Consistent with the scattered fluorescence observed in neuromasts after MTZ treatment (Fig. 1a), IBA1+ cells contained CHERRY fluorescent signals (Fig. 1g, Supplementary Fig. 2f and Supplementary Movie 2), which were likely derived from the engulfed CHERRY+ apoptotic cells or cell debris. Contrary to the neuromast, we found that, similar to the controls, the CHERRY fluorescence of the spinal cord remained unchanged after MTZ treatment in Sox2:Cherry-NTR axolotls (Fig. 1i, j). Overall, these results demonstrate effective ablation of Sox2+ cells in the body surface-located neuromasts, but not the CNS with MTZ treatment in Sox2:Cherry-NTR transgenic axolotls.

Effective ablation of the Sox2+ cells in the CNS with nifurpirinol in Sox2:Cherry-NTR knock-in axolotls
We then evaluated two nitro-heterocyclic derivatives, nifurpirinol (NFP) and ronidazole (RNZ), which have been reported as NTR substrates76,77, and identified that NFP is suitable candidate prodrug for NTR-mediated ablation in the axolotl (Supplementary Note 1). Specifically, NFP treatment induced progressive CHERRY signal decline across all Sox2-expression domains, including the noses, eyes, lateral line, spinal cord and brain in Sox2:Cherry-NTR axolotls, compared to that in DMSO and MTZ-treated animals (Supplementary Fig. 4a, b). By 20 dpt, spinal cord CHERRY signals became nearly undetectable (Supplementary Fig. 4a). Immunostaining on tail and brain sections revealed structural disruption and marked reduction of SOX2+ EGCs after NFP treatment (Fig. 2a, b, j), compared to MTZ-treated (Fig. 2c, k) or DMSO-treated controls (Fig. 2d, l). The number of the spinal cord EGCs decreased to 51.3% at 10 dpt (Fig. 2e), with no proliferative compensation in the residual SOX2+ population (Supplementary Fig. 5a–c), and a further reduction to 5.5% by 20 dpt (Fig. 2e). In contrast, the non-targeted tail PRRX1+ fibroblasts and PAX7+ muscle stem cells remained unchanged after NFP administration (Fig. 2f, g). Unexpectedly, we observed neuronal loss in both the spinal cord and the brain (Fig. 2b, g). In the spinal cord, NeuN+ or TUJ1+ neurons declined by approximately 50% at 20 dpt (Fig. 2h, i). This neuronal loss can be explained by the lineage relationship between EGCs and neurons, along with the presence of a SOX2+ NeuN+ neuronal subset (Fig. 2l, l’, l”). Moreover, co-expression of the cleaved Caspase-3 within the CNS SOX2+ cells confirmed that the NFP-induced ablation is mediated through apoptosis (Supplementary Fig. 5d–g). These results demonstrate that NFP is an effective prodrug for NTR-based targeted cell ablation, and achieves Sox2+ EGCs depletion via apoptosis in Sox2:Cherry-NTR axolotls.

Completely abolishing the CNS regeneration upon EGCs ablation
We next explored the role of EGCs in the CNS regeneration through EGC ablation using the optimized NFP administration protocol. Complete ablation of all Sox2+ cells in Sox2:Cherry-NTR axolotls caused lethality. To circumvent this, we transplanted spinal cords from Sox2:Cherry-NTR to d/d host to create chimeric axolotls. While spinal cord transplantation has been previously reported in axolotl, existing methods are intricate, labor-intensive, and inefficient23. We optimized the procedure by grafting donor spinal cord segments into the precisely anatomically matched vertebral canal of pre-amputated hosts (Fig. 3a). Following transplantation, the regenerated tail spinal cord exhibited CHERRY positivity (Fig. 3b), and all SOX2+ cells within it were CHERRY+ (Fig. 3c, d), indicating exclusive donor origin with no host contribution. Over 90% (28/30) of chimera contained proper integration of donor spinal cords (Supplementary Table 1), and it took less than 5 min to finish one transplantation. This established a streamlined spinal cord transplantation procedure, and we generated chimeric Sox2:Cherry-NTR axolotls for further functional analysis.

To investigate the roles of EGCs in spinal cord/tail regeneration, chimeric Sox2:Cherry-NTR axolotls were treated with NFP followed by tail amputation (Fig. 3a, e). Controls showed obvious tail regeneration, with CHERRY+ spinal cord extending into the tail outgrowth at 15 dpt and 20 dpt (Fig. 3f, g). In contrast, NFP treatment abolished CHERRY signal indicating depletion of Sox2+ EGCs (Fig. 3f, h) and completely blocked spinal cord regeneration (Fig. 3f, g). Moreover, tail regeneration failed, with fin tissue forming semicircular wound closure (Fig. 3f). At 20 dpt (10 dpa), immunostaining of section analysis revealed that NFP treatment ablated approximately 93% of SOX2+ cells in chimeric animals (Fig. 3h–j). Whereas 20% of SOX2+ cells were BrdU+ in controls, remaining SOX2+ cells in NFP-treated animals showed no BrdU incorporation (Fig. 3h–j). These results demonstrate that Sox2+ EGCs are essential for spinal cord regeneration and indirectly regulate tail regeneration.

Additionally, we observed a 35% reduction in NeuN+ neurons and a 23% reduction in TUJ1+ fluorescence intensity at 10 dpa compared with controls (Fig. 3k–p). Whole-mount immunostaining of regenerating tails from NFP-treated chimeras further showed both SOX2+ cells and TUJ1+ axons were sparse in the regenerated tail tip region, while control animals exhibited abundant SOX2+ cells in the spinal cord and TUJ1+ axons extending to the tail tip (Supplementary Fig. 6a, b and Supplementary Movie 3, 4). These findings collectively indicate that neuronal loss likely contributed to regeneration deficits in both the spinal cord and tail.

We further assessed the proliferation of host EGCs adjacent to the graft-host interface following the ablation of donor EGCs. Compared to controls (~16.5% BrdU+ in SOX2+ cells), the host EGCs exhibited a higher proliferating rate (~32.7%) in NFP-treated chimeras (Supplementary Fig. 6c–e). This localized proliferative response (>5 mm from amputation plane) is beyond the typical range of injury-induced activation (~500 μm)19, therefore, we considered this host EGCs proliferation induced by massive loss of donor EGCs rather than amputation.

Previous studies have suggested that EGCs may serve as a source for telencephalon regeneration in axolotls26–28. To assess EGC contribution to brain regeneration, similar to the approach applied in the spinal cord, we developed a telencephalon transplantation model in axolotls to generate chimeric Sox2:Cherry-NTR animals. The left telencephalons from Sox2:Cherry-NTR donors were transplanted into the cranial cavity of d/d hosts following removal of their original left telencephalon (Fig. 4a, b). This procedure enabled replacement of the host’s telencephalon with the donor tissue (Fig. 4a, c). Five months post-transplantation, CHERRY+ donor telencephalon integrated with d/d host brain, forming a chimeric architecture (50% of grafts maintaining CHERRY+ signal), with a transplantation success rate of approximately 16% (6/36) (Supplementary Table 2). Immunofluorescence in cross sections of chimeric brains revealed well integration of the transplanted telencephalon. SOX2+ EGCs lining the ventricular lumen expressed CHERRY (NTR) in donor telencephalon, whereas the host EGCs expressed only SOX2 (Fig. 4c). However, the donor telencephalon exhibited abnormal morphology with fragmented lumens (Fig. 4c). NFP treatment eliminated CHERRY signals in donor telencephalons of chimeric Sox2:Cherry-NTR axolotls by 45 dpt. CHERRY signals were undetectable, indicating ablation of Sox2+ EGCs, while the fluorescence in controls remained unchanged (Fig. 4d).

To investigate the brain regeneration in the absence of EGCs, we carried out brain injury on the transplanted left telencephalon of the chimeric Sox2:Cherry-NTR axolotls at 15 dpt, followed by BrdU pulse-chases at 10 and 20 days post-injury. Immunohistochemical analysis at 30 days post-injury demonstrated distinct regeneration outcomes (Fig. 4b, e–j). In the controls, we identified pronounced accumulation of SOX2+ EGCs, most of which were co-labeled with BrdU at lesions (Fig. 4f, h), indicating the involvement of EGCs in brain regeneration. Additionally, a substantial number of BrdU+ cells co-labeled with the neuronal subtype marker CTIP278, confirming the generation of new neurons after traumatic injury (Fig. 4j). In contrast, NFP-treated brains showed that along with drastic reduction of SOX2+ EGCs, both BrdU+ SOX2+ EGCs, and CTIP2+ BrdU+ neurons were near completely abolished, with evidence of fibrotic-like scarring (Fig. 4e, g, i).

In summary, EGC ablation via chimeric transplantation models blocks spinal cord and brain regeneration, establishing EGCs as the sole neuronal progenitors for CNS repair. Additionally, EGCs or spinal cord neuronal signals critically regulate tail regeneration.

Efficient ablation of cortical neurons with NTR 2.0 in the axolotls
Neuronal loss is the key feature of many CNS disorders. Restoration of lost neurons remains a great challenge. Therefore, establishing a neuronal ablation model is crucial to study the neuronal regenerative capacity. To this end, we generated NeuroD6:Cherry-NTR knock-in axolotls79 but found it challenging to ablate cortical neurons using the NTR/NFP system (Supplementary Note 2). To overcome this limitation, we next compared the original NTR with the recently published NTR 2.065 via brain electroporation80 and demonstrated that NTR 2.0 enables efficient neuronal ablation in axolotls when combined with MTZ (Supplementary Note 3), thus establishing the NTR 2.0/MTZ system as a superior method for neuronal ablation.

We then generated NeuroD6:Cherry-NTR 2.0 knock-in axolotls for targeted neuronal ablation (Fig. 5a–d). Live imaging revealed a strong increase of CHERRY fluorescence in the brains of NeuroD6:Cherry-NTR 2.0 larvae at 2 days post-MTZ treatment (Fig. 5e), likely originating from residual CHERRY signals of engulfed neuronal debris in microglia. The level of CHERRY gradually declined thereafter in axolotls treated with MTZ, whereas CHERRY expression in DMSO-treated controls remained constant (Fig. 5e). Immunohistochemical analysis of sections showed decreased Neurod6 mRNA and CHERRY expression (Fig. 5f), accompanied by massive depletion of cortical neurons (Fig. 5f–h), and a significant reduction in the thickness of the dorsal pallium, which was composed almost entirely of SOX2+ EGCs following neuronal ablation at 5 dpt (Fig. 5g, i). Furthermore, immunostaining revealed the co-localization of IBA1 and CHERRY at 5 dpt, indicating that microglia engulfed apoptotic neuronal debris. The persistent CHERRY+ signals likely originate from phagocytosed neuronal debris within microglia that has not been completely degraded (Fig. 5e, j). Behavioral assays demonstrated that upon ablation of the nearly entire population of dorsal pallium neurons, axolotls failed to swallow the artemia food (Supplementary Fig. 9a, b and Supplementary Movie 5, 6). At 10 dpt, CHERRY+ cells started to repopulate the pallium, along with the onset of NeuroD6 mRNA expression determined by in situ hybridization, and eventually restored by 20 dpt (Fig. 5f). With completion of brain regeneration, the food-swallowing defects were functionally recovered (Supplementary Fig. 9c, d and Supplementary Movie 7, 8). These results conclusively demonstrate that the NTR 2.0/MTZ system achieves robust and specific neuronal ablation in axolotls.

In adult axolotl pallium, the NeuroD6+ neurons distributed across the cortex co-express diverse neuronal subtype markers26,28,81,82 (Supplementary Fig. 10b). To investigate the spatial organization and subtypes of regenerated neurons after large-scale neuronal ablation in the dorsal pallium, and to avoid developmental processes confounding regenerative responses, we carried out neuronal ablation in adult (15 cm in size from snout to tail tip) NeuroD6:Cherry-NTR 2.0 axolotls and analyzed the neuronal regeneration (Supplementary Fig. 10a). Immunohistochemical analysis showed that, similar to phenotypes observed in larvae, the thickness of the cortex is markedly reduced to about 200 μm by 10 days post-MTZ treatment in NeuroD6:Cherry-NTR 2.0 axolotls, compared to 400 μm in the controls (Supplementary Fig. 10c–i), accompanied by a significant reduction of CHERRY signals (Supplementary Fig. 10c–h). The CHERRY+ NEUN+ cortical neurons, including CHERRY+ CALB2+, and CHERRY+ SATB1+ neuronal subtypes were nearly completely lost in NeuroD6:Cherry-NTR 2.0 axolotls by 10 days post-MTZ treatment, compared to that in the controls (Supplementary Fig. 10c–h, j–l). With a single pulse of BrdU administration, we observed an increased number of SOX2+ BrdU+ cells in the dorsal pallium in NeuroD6:Cherry-NTR 2.0 axolotls at 10 dpt, compared to the controls (Supplementary Fig. 10m–o), suggesting elevated proliferation of EGCs after cortical neuron ablation.

To track and distinguish the early and later born neurons after large-scale cortical neuron ablation, we performed birth-dating experiment via sequential BrdU and EdU administrations, then analyzed the regenerated cortex at 113 dpt (Fig. 6a). Immunohistochemical analysis revealed that the sequentially labeled regenerated neurons formed a laminar-like distribution in the dorsal pallium, where early born neurons (BrdU+ SOX2−) were predominantly localized to superficial zones, while later born neurons (EdU+ SOX2−) preferentially settled in deeper zones (Fig. 6b, c). In species such as frogs and chicks, it has been reported that the organization of the cortical neurons follows a chronological order during pallium development, with an “outside-in” pattern: early-born neurons occupying superficial zones and late-born neurons residing deeper zones83–87. We carried out a similar birth-dating experiment in d/d axolotl larvae (Fig. 6d) and found that during development, cortical neurons are organized as “outside-in” pattern similar to that identified in regeneration (Fig. 6e, f). Next, we examined whether all cortical neuron subtypes were regenerated or not after large-scale cortical neuron ablation. By 113 dpt, immunohistochemical and mRNA in situ hybridization analysis demonstrated that pan neurons labeled by NEUN, and dorsal-located CALB2+, SATB1+, Tmie+, Tbr1+, Nptx+, and Fezf2+ neuronal subtypes were restored and organized as that in the controls (Fig. 6g–y and Supplementary Fig. 11a–d). A subpopulation of NEUN+, CALB2+, and SATB1+ neurons was incorporated with BrdU (Fig. 6j, p, v), indicating de novo regeneration of ablated neurons or neuronal subtypes. Furthermore, measuring the cortical thickness showed that the partial recovery of the cortex, likely due to the insufficient regeneration duration (Fig. 6z). These data collectively demonstrate the recovery of the cortical neurons and neuron subtypes and their organization after nearly entire cortical neuron ablation in axolotl.

Generation of a versatile Cre inducible NTR 2.0 transgenic line
Generating a stable knock-in NTR 2.0 transgenic line is typically time- and labor-intensive. In axolotls, many Cre and inducible Cre transgenic lines have been generated. To expand the application of NTR 2.0/MTZ system, we developed a CAGGs:loxP-BFPnls-loxP-Cherry-P2A-NTR 2.0 transgenic axolotl line for Cre-loxP-based conditional targeted cell ablation (Fig. 7a). When crossed with a specific Cre transgenic line, the floxed stop cassette is excised, resulting in Cherry-NTR 2.0 expression under the control of the constitutive CAGGs promoter, thereby enabling targeted ablation of specific cells. In the established CAGGs:loxP-BFPnls-loxP-Cherry-P2A-NTR 2.0 line, we observed ubiquitous BFP expression throughout the body (Fig. 7b). However, in line with previous observations, the BFP fluorescence signal in muscle was stronger than that in the CNS and the fin tissue38,88,89. Before conversion, CHERRY fluorescence was absent across the body, except for spontaneous fluorescence in the abdominal region (Fig. 7b) To test the Cre-mediated NTR 2.0 conversion and targeted cell ablation, we introduced pCAGGs:Cre plasmid into the spinal cord and muscle tissue of the tail in CAGGs:loxP-BFPnls-loxP-Cherry-P2A-NTR 2.0 axolotl, followed by electroporation (Fig. 7c and Supplementary Fig. 12a). We observed robust CHERRY fluorescence expression at 3 days post-electroporation (Fig. 7d and Supplementary Fig. 12b), indicating efficient Cre-mediated conversion. Subsequent MTZ treatment of electroporated animals led to significant attenuation of the CHERRY signal compared to that in DMSO-treated controls (Fig. 7d, e and Supplementary Fig. 12b). Immunofluorescence analysis of spinal cord sections revealed absence of CHERRY signals and collapse of the spinal cord structure after MTZ treatment, compared to the controls in which CHERRY were clearly expressed in the intact spinal cord (Fig. 7f). These results demonstrates that targeted cell ablation mediated by exogenous Cre is achievable in the CAGGs:loxP-BFPnls-loxP-Cherry-P2A-NTR 2.0 line.

We next combined the CAGGs:loxP-BFPnls-loxP-Cherry-P2A-NTR 2.0 axolotls with an inducible Cre transgenic line to evaluate the cell ablation efficiency. We bred CAGGs:loxP-BFPnls-loxP-Cherry-P2A-NTR 2.0 with the inducible Nkx2.2:memCherry-T2A-Cre-ERT2 line75 to create CAGGs:loxP-BFPnls-loxP-Cherry-P2A-NTR 2.0/Nkx2.2:memCherry-T2A-Cre-ERT2 double transgenic axolotls (dTG) (Fig. 7g). Nkx2.2 is expressed in ventral EGCs in the midbrain. Prior to tamoxifen treatment, we observed a dim membrane-tagged CHERRY expression in the midbrain driven by the endogenous Nkx2.2 promoter (Fig. 7i). Upon tamoxifen administration, Cre-mediated removal of the floxed stop cassette led to the CHERRY-NTR 2.0 expression under the control of the CAGGs promoter in Nkx2.2+ cells (Fig. 7g). At 8 days post tamoxifen treatment, we found significantly elevated CHERRY expression in the midbrain in tamoxifen-treated animals compared to untreated controls (Fig. 7h, j). Immunohistochemical analysis revealed the switch from a dim membrane-tagged CHERRY to strong cytosolic CHERRY signals in a subpopulation of NKX2.2+ cells, indicating the conversion and expression of NTR 2.0 in these cells (Supplementary Fig. 12c). Subsequent MTZ treatment resulted in a remarkable reduction of CHERRY fluorescence at 6 days post-MTZ treatment (Fig. 7j). Immunofluorescence staining showed that the loss of cytosolic CHERRY-expressing cells in the midbrain in MTZ treated dTG axolotls compared to that in the controls, suggesting specific ablation of Cre-converted NKX2.2+ cells (Fig. 7k and Supplementary Fig. 12d). Overall, these results demonstrate the Cre-LoxP-based conditional NTR 2.0 transgenic line we have generated enables tissue-specific ablation of the targeted cells when combined with established Cre line.

Methods
Ethical statement
Axolotls (A. mexicanum) were housed and bred in dechlorinated tap water at 18°C–20°C with a 12-h light/12-h dark cycle. Before performing experiments such as electroporation, transplantation, live imaging and harvest, animals were deeply anesthetized using 0.03% benzocaine (Sigma-Aldrich, E1501). In this study, animal care was in accordance with the animal welfare legislation in China, and all animal experiments were approved by the Biomedical Research Ethics Committee of Guangdong Provincial People Hospital (license number: KY-Q-2022-395-03).

Animals
The d/d axolotl stain of A. mexicanum, which was used for breeding to generate transgenic axolotls by single-cell stage egg injection, was originated from Ambystoma Genetic Stock Center (https://ambystoma.uky.edu/genetic-stock-center/). The Nkx2.2:memCherry-T2A-Cre-ERT2 transgenic line has been previously described75. Sox2:Cherry-NTR, NeuroD6:Cherry-NTR, NeuroD6:Cherry-NTR 2.0 and CAGGs:loxp-BFPnls-loxp-Cherry-P2A-NTR 2.0 transgenic lines were generated in this study.

No sex or gender-specific bias was incorporated into the experimental design. This study utilized axolotls of three distinct size classes (body-length from snout to tail tip): 1.5–3 cm, 4–8 cm, and 15–18 cm at the time of prodrug administration. Among these developmental stages, only the largest size (>15 cm) permitted morphological sex determination, whereas smaller individuals would require genotyping for accurate sex identification. Animals were randomly assigned to experimental groups without sex identification. Experimental conditions were not stratified by sex, and the observed biological phenomena are presumed to be sex-independent. Consequently, no sex-based analyses were performed.

The Sox2-gRNAs and NeuroD6-gRNAs used for generating transgenic axolotls were previously described75 and were synthesized by Genscript in this study. The gRNA targeting sites are listed below:
Sox2-gRNA#1 (5′-ATGTGACAATGAGCGCGG-3′),
Sox2-gRNA#2 (5′-CTCATTGTCACATGTGCGAG-3′),
Sox2-gRNA#3 (5′-TGGCGAGAGTCTGTCTGTGC-3′),
NeuroD6-gRNA#1 (5′-ACAAAGCAGATGCTAGTTGA-3′),
NeuroD6-gRNA#2 (5′-AGTTGACGGAGTAATTGACA-3).

Plasmids construction and gRNAs
All plasmids were constructed using restriction enzyme-based cloning and Gibson assembly, followed by Sanger sequencing verification (Tsingke, China). The primers used for plasmids construction were listed in Supplementary Table 3. Plasmid purification was performed with the EndoFree Plasmid Kit (12162, Qiagen, Germany) for further experiments.

(1) For construction of the in vivo electroporation plasmids pCAGGs:Cherry-NTR, pCAGGs:Cherry-NTR 2.0 and pCAGGs:Cre, the Cherry-NTR63 and Cherry-P2A-NTR 2.065 coding sequences were commercially synthesized (Genscript, China), and the Cre coding sequence was amplified from the previously described pGEMT-Nkx2.2-P2A-memCherry-T2A-Cre-ERT2-PA plasmid75. The Cherry-NTR 2.0 fragment was constructed by PCR amplification using Gibson assembly primers (listed in Supplementary Table 3). The Cherry-NTR, Cherry-NTR 2.0 and Cre fragment were then subcloned into pCAGGs vector using Gibson Assembly Master Mix kit (E2611, NEB, USA) according to the manufacturer’s instruction.

(2) To prepare targeting construct pGEMT-Sox2-CDS-T2A-Cherry-NTR-pA that was used for generating the Sox2: Cherry-NTR knock-in line, the previously described pGEMT-Sox2-CDS-T2A-Cherry-pA plasmid75 was linearized and excised mCherry sequence using restriction enzymes BmgBI and EcoRI. The Cherry-NTR fragment was subsequently inserted into the linearized vector according to manufacturer’s instruction of Gibson assembly kit.

(3) To prepare targeting construct pGEMT-NeuroD6-CDS-T2A-Cherry-NTR-pA that was used for generating the NeuroD6:Cherry-NTR knock-in line, the previously described pGEMT-Sox2-CDS-T2A-Cherry-NTR-pA plasmid was linearized and excised Sox2 CDS sequence using restriction enzymes MluI and SphI. The NeuroD6 gene was identified and validated in the axolotl genome (v6.0-DD assembly). The NeuroD6-CDS fragment was amplified from axolotl genomic DNA and was subsequently inserted into the linearized vector according to manufacturer’s instruction of Gibson assembly kit.

(4) To prepare the targeting construct pGEMT-NeuroD6-CDS-T2A-Cherry-NTR 2.0-pA that was used for generating the NeuroD6:Cherry-NTR knock-in line, pGEMT-Neurod6-CDS-T2A-Cherry-NTR-pA plasmid was linearized and excised NTR sequence using restriction enzymes BsrGI and BamHI. The NTR 2.0 fragment was subsequently inserted into the linearized vector according to manufacturer’s instruction of Gibson assembly kit.

(5) To prepare the targeting construct pCAGGs: loxp-BFPnls-STOP-loxp-Cherry-P2A-NTR 2.0, the previously described pCAGGs: loxp-BFPnls-STOP-loxp-Smad7-Cherry94 plasmid was linearized and excised Smad7-Cherry sequence using restriction enzymes XmaI and ClaI. The Cherry-P2A-NTR 2.0 fragment was subsequently inserted into the linearized vector according to manufacturer’s instruction of Gibson assembly kit.

Generation of transgenic axolotls
To generate transgenic axolotls, the mixture preparation of construct-gRNAs-CAS9-NLS RNP complexes or construct-Tol2 mRNA and egg injection were performed following established protocol95,96. The construct-gRNAs-CAS9-NLS RNP complexes were composed of 50 ng/µL targeting construct, 0.5 µg/µL CAS9-NLS protein, and 0.4 µg/µL gRNA in 1× CAS9 buffer. The construct-Tol2 mRNA mixture contained 20 ng/µL targeting construct and 75 ng/µL Tol2 mRNA in nuclease-free water. For egg injection, freshly laid single-cell stage axolotl eggs were collected and dejellied in 1× MMR solution, and then transferred to 1× MMR with 10% Ficoll for injection. The construct-gRNAs-CAS9-NLS RNP complexes or construct-Tol2 mRNA mixture was loaded into a microinjection glass capillary and injected into single-cell stage eggs (5 nL per egg) with the assistance of Pneumatic PicoPump (WPI, PV830, USA) and a stereoscope. Injected eggs were then incubated in 1× MMR with 10% Ficoll for 2–3 h, followed by overnight incubation in 0.1× MMR with 5% Ficoll. Before developing to the gastrula stage, the eggs were transferred to 24-well plates containing 0.1× MMR and raised there until the hatching stage. Transgenic axolotls were confirmed by both phenotyping and genotyping. For phenotyping, the larvae were screened using the fluorescence stereoscope (Olympus, SZX16, Japan) to detect reporter fluorescence in specific tissues and organs. For genotyping, the alkaline lysis method was employed for genomic DNA extraction95, followed by genotyping PCR with primers listed in Supplementary Table 3.

Axolotl spinal cord, brain and muscle electroporation
Electroporation of axolotl spinal cord and brain tissues was conducted as previously described80,97,98. Briefly, plasmids (pCAGGs:Cherry-NTR, pCAGGs:Cherry-NTR 2.0 and pCAGGs:Cre) were diluted to a 2-3 µg/µL in 0.8× PBS containing 0.05% Fast Green dye for visual tracking. After the animals were anesthetized with 0.03% benzocaine, plasmid solutions were microinjected into either the central canal (for the spinal cord, 0.2–0.3 µL per animal) or lateral ventricle (for the brain, 2 µL per animal) using calibrated glass micropipettes. Immediately post-injection, electroporation was performed in ice-cold PBS with an electroporator (Nepagene, NEPA21, Japan) equipped with 10-mm paddle electrodes (Nepagene, CUY650P10, Japan), applying distinct parameter sets: for spinal cord delivery, 70 V poring pulse (5 ms) followed by four 40 V bipolar transfer pulses (50 ms) at 999 ms intervals with 10% decay; for brain delivery, 80 V poring pulse (5 ms) preceding five 60 V unipolar transfer pulses (50 ms) under equivalent interval/decay settings as those for the spinal cord. After the procedure, the animals were returned to dechlorinated tap water to recover.

For axolotl muscle electroporation, pCAGGs:Cre plasmid was diluted to 2–3 µg/µL in 0.8× PBS with 0.05% Fast Green dye. The solution was subcutaneously injected into the lateral region of the tail (1–2 µL per animal). Electroporation was conducted in ice-cold PBS with the same parameter sets as those employed for spinal cord delivery.

Nitroreductase substrate treatment
Nitroreductase substrates metronidazole (Sigma-Aldrich, M1547), nifurpirinol (Dr Ehrenstorfer, C15523050), and ronidazole (Sigma-Aldrich, R7635) were dissolved in dimethyl sulfoxide (DMSO, Sigma-Aldrich, D8418) to generate stock solutions (DMSO final concentration ≤0.2% in all treatment groups, with 0.2% DMSO alone serving as the vehicle control). Animals were exposed to the substrates via full-immersion bath treatments in dechlorinated tap water. Dose-response experiments tested the following concentration ranges: metronidazole (1.25–40 mM), nifurpirinol (1.25–40 µM), and ronidazole (0.25–8 mM). To prevent photodegradation of the nitroreductase substrates, animals were maintained under light-protected conditions throughout the treatment period.

4-hydroxytamoxifen treatment
4-hydroxytamoxifen (4-OHT) was prepared as a 20 mM stock solution in DMSO, and diluted to generate a working solution in dechlorinated tap water. To activate the Cre-ERT2 system, larvae (1.5 cm body length from snout to tail tip) were treated with 2 µM 4-OHT through eight sequential 12-h exposures, each separated by a 12-h interval.

Axolotl spinal cord transplantation
We used donor Sox2:Cherry-NTR and host d/d axolotls (4 cm in body length) for spinal cord transplantation. All animals were deeply anesthetized prior to surgery. Both donor and host animals were amputated at the equivalent somite level (12–14th somite rostral to the cloaca) to expose the amputation plane. For the host, A lateral incision was made through the skin at 2 somites rostral to the amputation plane using fine surgical scissors, and the spinal cord was transected at this site. A 2-somite-length spinal cord segment was then carefully extracted from the spinal canal through the amputation plane using sterile forceps, with gentle handling to preserve structural integrity.

For the donor, A 3-somite-length spinal cord segment at the equivalent anatomical site was removed using the same extraction protocol: a lateral skin incision near the amputation plane, transection of the spinal cord, and extraction of the target segment from the spinal canal via the amputation plane. After extraction, the damaged end of the donor segment was slightly trimmed to refine the graft for transplantation. Immediately following host spinal cord removal, the donor segment (trimmed to match the 2-somite-length of the host’s excised segment) was inserted into the host’s resulting spinal canal cavity. This ensured precise anatomical alignment and direct contact between the transplanted graft and the host’s spinal cord stumps. Two months after spinal cord transplantation (at 7 cm in body length), the success of spinal cord transplantation was defined by normal morphology in the regenerated tail segment, in which presented the donor-derived CHERRY positive spinal cord.

Axolotl telencephalon transplantation and brain injury
Donor Sox2:Cherry-NTR and host d/d axolotls (5–6 cm body length) were utilized for telencephalon transplantation experiments. Prior to surgery, all animals were deeply anesthetized. For telencephalon transplantation, a scalp flap and cranial flap were created on the top of the animals’ head to expose the left telencephalon. The host’s left telencephalon and olfactory bulb were microsurgically excised by transecting the olfactory nerve root and telencephalon-diencephalon boundary with spring scissors (Fine Science Tools, 15003-08, USA). Donor tissue (left telencephalon with intact olfactory bulb) was harvested using identical methods and transplanted into the host’s cranial cavity. The graft was positioned to ensure anatomical alignment between donor telencephalic margins and host diencephalic edges. Subsequently, the cranial flap and scalp flap were repositioned to facilitate recovery. Five months after brain transplantation (reaching 17–18 cm in body length), the success of telencephalon transplantation was defined by normal morphology in the integrated telencephalon, in which presented the donor-derived CHERRY positive telencephalon.

Axolotl brain injury was conducted as previously described28. Briefly, deeply anesthetized animals were made a rectangular cranial skin/skull flap over the left telencephalon, using a scalpel and spring scissors and reflected to expose the telencephalon. For the injury, a 0.5 mm × 0.5 mm square of tissue was excised from the dorsal region of the left telencephalon. To standardize the lesion site across animals, the excision was precisely localized between the choroid plexus and olfactory bulb of the left telencephalon. After injury, the cranial skin/skull flaps were repositioned without sutures.

BrdU and EdU injection and detection
Anesthetized animals received intraperitoneal injections of BrdU (Sigma-Aldrich, B9285) or EdU (Thermo Scientific, A10044). The required injection volume for BrdU (10 mg/mL) or EdU (1 mg/mL) was calculated for each animal based on its individual body weight, ensuring precise delivery of the specified doses. For BrdU administration, 50 µg/g body weight for single-pulse labeling or 25 µg/g body weight per dose for multi-dose regimens. For EdU administration, 10 µg/g body weight for each pulse.

For BrdU immunodetection, cryosections were treated with 2 M HCl at 37°C for 10 min to denature the DNA. These sections were then washed five times with 1× PBS for 20 min each before immunofluorescence staining was performed following the immunofluorescence staining protocol.

EdU incorporation was visualized using Click-iT EdU (Thermo Scientific, C10634) following the manufacturer’s protocol. This mild labeling protocol preserved epitope integrity, allowing subsequent immunofluorescence staining according to the immunofluorescence staining protocol.

Samples collection and cryosection
The harvested samples were fixed in MOPS/EGTA/magnesium Sulfate/4% formaldehyde Buffer (MEMFA) at 4°C with the following durations: overnight for tails and 2–3 days for brains. After fixation, specimens were dehydrated in 30% sucrose in 1× PBS at 4°C and then snap-froze and embedded in OCT compound (Sakura, 4583, USA). The embedded blocks were stored at −80°C until subsequent processing. Serial sections were obtained using a cryostat (Thermo Scientific, Cryostar NX70, USA) with optimized section thickness: 10-µm coronal tail sections and 16-µm coronal brain sections. The cryosections were archived at −20°C until histological staining and analysis.

RNA probe and mRNA in situ hybridization
In situ hybridization probes were designed based on the axolotl transcriptome assembly v4.7 (https://www.axolotl-omics.org/)99. Gene fragments were PCR-amplified from axolotl brain cDNA using gene-specific primers (listed in Supplementary Table S3). The amplified products were sequence-verified, and then used as templates in a secondary PCR with reverse primers containing T7 promoter sequences (5′-TAATACGACTCACTATAGGG-3′) to generate templates for in vitro transcription. Digoxigenin-labeled antisense RNA probes were synthesized via in vitro transcription using the DIG RNA labeling mix (Roche, 11277073910). For in situ hybridization, 16-µm-thick coronal brain cryosections were hybridized with 500 ng/mL DIG probes at 60°C overnight, followed by immunological detection using alkaline phosphatase-conjugated anti-DIG Fab fragments (1:5000, Roche, 11093274910) and chromogenic development with BM purple substrate (Roche, 11442074001) as previously described38.

Immunofluorescence staining on sections
The immunofluorescence staining was performed as using a standard protocol89. Briefly, sections were washed with 1× PBS/0.3% Triton X-100 and blocked in 5% donkey serum prepared in 1× PBS/0.3% Triton X-100 for 2 h at room temperature. Sections were then incubated overnight at 4°C with primary antibodies diluted in commercial antibody diluent (Genefist, GF-1600-01). Following incubation, sections were washed extensively with 1× PBS/0.3% Triton X-100, and incubated with secondary antibodies with proper dilution in antibody diluent for 2 h at room temperature.

The following primary antibodies were used against the corresponding antigens: SOX2 (1:500, Abcam, ab97959; 1:200, Abcam, 14-9811-82), NeuN (1:300, Sigma, MAB377), TUJ1 (1:500, R&D, MAB1195), Cleaved Caspase-3 (1:500, CST, 9661s), BrdU (1:500, Invitrogen, MA3-071; 1:500, Abcam, ab1893), CTIP2 (1:500, Abcam, ab18465), CALB2 (1:500, Swant, 7697), SATB1 (1:20, Abcam, ab51502), Iba1 (1:500, Wako, 019-19741), CHERRY (1:1000, Invitrogen, M11217), NKX2.2 (1:100, BD Biosciences, 564731). Fluorescent labeling was performed using secondary antibodies conjugated to Cy3, Alexa 488, Alexa 647, or Alexa 750 (1:500 dilution). Nuclei were counterstained with DAPI (1:1000, Sigma, D9542).

Neuromasts whole-mount immunofluorescence staining
The MEMFA-fixed tail samples were rinse in PBS at room temperature (RT) for 3×30 min. After washing, the tail skin was carefully dissected to remove attached muscle tissue by forceps. The isolated skins were dehydrated through an ascending methanol/H2O gradient (25%, 50%, 80%, 100%; 1 h per step at RT), followed by an additional 1-h wash in 100% methanol and chilling at 4°C. Tissues were incubated for 3 h at RT in 66% dichloromethane (DCM; Sigma 270997)/33% methanol, then washed twice in 100% methanol, and chilled again at 4°C. The samples were then bleached overnight at 4°C in freshly prepared 5% H2O2 in methanol. Following the bleaching step, the tissues were rehydrated through a descending methanol/H2O gradient (80%, 50%, 25%, and finally PBS; 1 h per step at RT). After rehydration, tissues were washed in PBS containing 0.3% Triton X-100 for 3×15 min, and blocked in PBS with 0.3% Triton X-100 and 10% donkey serum for 2 h at RT. Primary antibody against SOX2, CHERRY and IBA1 were diluted in antibody diluent, the incubation was carried out overnight at 4°C, followed by washes in PBS with 0.1% Tween-20 (PBS-Tw) for 5×10 min. Secondary antibody staining was performed overnight at 37°C, followed by PBS-Tw washes for 5×10 min at RT.

The skin samples were stored in PBS-Tw at 4°C. Neuromasts were imaged on the laser scanning confocal microscope (Zeiss, LSM980, Germany) using a 20× objective and Z-stack acquisition. Image visualization and quantification were performed using Imaris 9.9.1.

Whole-mount iDISCO clearing
Tail tissue staining clearing and immunostaining were performed according to the iDISCO protocol100,101. Briefly, MEMFA-fixed tail samples were rinse in PBS, then dehydrated through an ascending methanol gradient (20, 40, 60, 80, and 100%). After 100% methanol incubation at 4°C, tissues were treated overnight at RT with 66% DCM and 33% methanol. Samples were then washed twice in 100% methanol, chilled at 4°C, and bleached in freshly prepared 5% H2O2 in methanol overnight at 4°C. Following bleaching, rehydration was performed using a descending methanol gradient (80, 60, 40, and 20%) at RT. Rehydrated samples were washed twice in PBS containing 0.2% Triton X-100. Tissues were permeabilized for 24 h at 37°C in a solution of 0.2% Triton X-100, 20% DMSO, and 0.3 M glycine.

For immunostaining, tails were blocked with 5% donkey serum in PBS containing 0.2% Triton X-100 and 10% DMSO, then incubated with primary antibody against SOX2 and TUJ1 diluted in antibody diluent at 37°C for 3 days. After primary antibody incubation, tissues were washed 4–5 times over 24 h in PBS containing 0.2% Tween-20 and 10 mg/L heparin (PBTwH). The samples were then incubated with the secondary antibody in antibody diluent at 37°C for 3 days, followed by PBTwH washes.

For clearing, samples were embedded in 1% agarose and dehydrated through a methanol gradient. Tissues were incubated for 3 h at RT with shaking in 66% DCM/33% methanol, washed twice in 100% DCM, and finally transferred to dibenzyl ether (DBE; Sigma, 108014-1KG) for clearing.

The clearing samples were imaged with a Lightsheet microscope (Zeiss, Lightsheet 7) at 20× magnification and 0.56 µm resolution. The acquired images were subsequently visualized and analyzed using Imaris 9.9.1.

Imaging
Live brightfield and fluorescence imaging of axolotls was performed using a stereo fluorescence microscope (Olympus, SZX16). In situ hybridization signals in cryosections were imaged with an inverted microscope (Olympus, IX83).

For immunostaining analysis, fluorescence imaging was conducted using either the IX83 inverted microscope system or a laser scanning confocal microscope (Zeiss, LSM980, Germany). The whole-mount clearing samples were imaged with a Lightsheet microscope (Zeiss, Lightsheet 7).

Statistics and reproducibility
Statistical analyses were conducted using GraphPad Prism software (Version 10.3.0, GraphPad Software, USA). Data are presented as mean ± standard error of the mean (SEM) from at least three independent biological replicates. Comparisons between two independent groups were analyzed using an unpaired two-tailed Student’s t-test. The sample sizes (n) and relevant quantification conditions were reported in the corresponding figure legends. No statistical method was used to predetermine sample size; no data were excluded from the analyses. Animals were randomly assigned to groups for experimental analyses. All experiments are independently repeated at least two times with similar results. A p-value less than 0.05 was considered statistically significant. The quantitative data used in this study are provided in the Source data file. For graphical representations, n.s indicates P > 0.05, * indicates P < 0.05, ** indicates P < 0.01, *** indicates P < 0.001, **** indicates P < 0.0001.

Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability
All data supporting the findings of this study are available within the article and its Supplementary Information files or from the corresponding author upon request. The plasmids and transgenic axolotl lines generated in this study are available to the research community upon request. The quantitative data used in this study are provided in the Source Data file. Source data are provided with this paper.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Cell ablation tools are currently not available for studies using the axolotl, which arguably is the best tetrapod model for\nregeneration research. The manuscript chronicles a series of experiments that establish efficacy in using NTR-NTR2.0/MTZ\ncell ablation to target neuronal cell types in the lateral line, spinal cord, and brain. Additionally, the study reports\ndevelopment of a Cre-loxP-based reporter NTR2.0 line that should allow targeted ablation of any axolotl Cre line. Although\nthe manuscript is methodologically oriented, several interesting biological findings are reported to establish broader\nsignificance of the work, including Sox2+cell ablation that inhibited spinal cord and tail regeneration, and Neurod6+ cell\nablation that provided an opportunity to detail neuronal patterning in the brain of a regeneration competent model.\nImpressive amount of work accomplished using the axolotl. I list some minor issues below.\nLine 40: I think \"chronologically\" is the correct word, and can you please describe more precisely what is meant by layer\norganization.\nLine 41-42: Perhaps change the sentence to this: \", enabling tissue-specific ablation of specific cell types when crossed to\nexisting Cre lines.\nLine 54: Maybe change to \"knowledge gaps hinder\".\nLine 61: Maybe change to \"using genetic approaches\".\nLine 79: Italicize \"E. coli\".\nLine 88-89: Please rewrite this sentence: \"While axolotls exhibit remarkable brain regeneration after trauma, therefore it\nserves as an ideal model for brain regeneration research.\"\nLines 89-91: Many studies predating single cell sequencing established a primary role for EGCs in spinal regeneration and\nprobably should be referenced.\nLines 127-129: Maybe rewrite these two sentences as they do not read well.\nLines 139, 217, 221: remove comma.\nLine 140: delete \" that from the\".\nLine 149: delete \"live imaging\".\nLine 151: change \"the identified macrophages\" to IBA1+ cells.\nLines 171 and 180: I think \"compared\" is the correct word.\nLine 232: maybe change \"on\" to in.\nLine 234. Maybe end sentence with a period after telencephalon, then start next sentence with Sox2+.\nLine 238. Maybe end sentence with a period after 45 dpt, then start next sentence with CHERRY signals were undetectable,\nindicating ablation of Sox2+ EGCs, while fluorescence in\ncontrols remained unchanged.\nLine 247: remove comma after BrdU.\nLine 249: include a reference establish CTIP1 as a marker of new born neurons.\nLine 252: maybe change \"wounds healing via\" with evidence of.\nLine 257: maybe change \"neurons-derived\" to neuronal.\nLine 262: It is debatable that relatively rapid depletion of cortical neurons by cell ablation provides a model for protracted cell\nloss seen im many degenerative neuronal disorders. Maybe clarify/better rationalize the manipulation as a model.\nLine 268: change \"on\" to of.\nLine 289: change sentence maybe to this: These results established NTR2.0/MTZ as superior method for neuronal ablation.\nLine 294: change \"originated\" to originating.\nLine 297: change \"on\" to of.\nLine 300: please rewrite \" that comprised near solely\".\nLine 313: please clarify what is meant by the developmental effect. Also, were behavioral changes observed in adult axolotls\nas reported in larval axolotls.\nLine 322: change to nearly completely lost.\nLine 358-360. Please rewrite this sentence to achieve clarity.\nLine 373: change \"on\" to of.\nLine 399: change \"enable\" to enables.\nLine 408: I don't think the results of the study enabled modeling of neurodegenerative disease, maybe tone this statement\ndown.\nLines 438-449: Maybe good to cite again/comment on the more recent approach KID/KIS as this mediates cell-ablation more\nrapidly in zebrafish embryos than NTR2.0.\nLines 455-458: Please rewrite this sentence to achieve clarity, and perhaps cite relevant research supporting the idea of low\nCre conversion rate.\nLines 460-466: There are several small grammatical errors in this paragraph, for example on line 464, maybe change \"\ntargeted cells ablation are\" to \"cell ablation is\".\nLines 482-484: How will the lines created in the study be made available for use by other researchers? Will they be donated\nto a stock center? This would seem important to increase axolotl research efforts community-wide.\nLines 546 and 563 and 574 and 631: What volume amounts were injected?\nLine 600: How exactly was the spinal cord removed? Can more detail be provided?", "reviewer_2": "In this manuscript, Fu et al develop the nitroreductase system for ablation of glial cells and neurons in the central nervous\nsystem in axolotls. This is a widely used system in zebrafish and mice, and could, in theory could be a very useful system to\nadapt in axolotls to ablate certain cell populations specifically. The manuscript is long, showing a lot of things that did not\nwork efficiently. Overall, the manuscript could be shortened and transformed into a more concise Methods paper. However, I\nhave some major concerns about the ablation efficacy shown in the manuscript. Specific points:\n1. The authors first test a ubiquitous promoter CAGG driving Cherry NTR and quantify fluorescence intensity in low mag\nimages and show a decrease at 6 days post-treatment. This does not prove that these cells types are actually ablated. I\nwould remove these panel Figure 1a-e and the text.\n2. The authors then generated a Sox2-Cherry-NTR line, these panels could be moved to Supplementary data.\n3. Figure 2 is confusing /unconvincing. Panel A shows that at 2 and 5 days post-treatment with MTZ, there is lower\nfluorescence in the lateral line cells, but it is still detected. But Panel C at 3dpt appears to show almost no Sox2+ cells after 3\ndays treatment, however, the area of skin shown is much thinner than control, also like it also doesn’t have any lateral line\ncells here. This result needs quantification, whereby number of cells per neuromast that are expressing Sox2 are counted\nprior to giving MTZ, then the number of Sox2 positive cells counted 1dpt, 2dpt, etc. Number of animals used needs per\nexperimental condition needs to be stated in the Figure legend.\n4. Figure 3 the authors determine that NFP is an effective drug for ablation of Sox2 glial cells in the spinal cord. However it is\nvery concerning that it takes 10 days of treatment for 51 of the cells to be ablated. When 50% of cells are ablated do the\nsurrounding glial cells begin to proliferate to compensate for the dying cells?\n5. The fact that the authors see 50% decline in spinal cord neurons at 20dpt is not surprising since these animals are still\ngrowing and the glial cells give rise to new neurons and sox2 is a neural progenitor marker.\n6. In Figure 4 If they administer NTR for 20 days to ablate the Sox2 positive cells in the spinal cord, then the animals die, this\noverall suggests that bathing the animals in the drug for 20 days maybe the cause of death and not the ablation of the cells.\nIn the test of nonspecific toxicity of the drug they only tested to 10 days post treatment, a control for 20 dpt needs to be done.\nHave you tried injecting the drug into the lumen of the spinal cord instead of bathing the animals?\n7. In Figure 4 the authors embark on a complicated unnecessary experiment to show that glial cells are necessary for spinal\ncord regeneration. The authors create chimeric animals by spinal cord transplantation. The better experiment would be to\nsimply treat for 5 days with NTR, amputate the tail, continue treatment for 5days, whereby at least 50% of the cells should be\nablated and measure the length of the regenerate overtime versus the control animals.\n8. Line 219, referring to Figure 4, “ neuronal loss likely contributed to this phenotype”. The authors need to quantify neuronal\nregeneration in control versus ablated animals by carrying out NeuN staining on sections and quantifying and doing a\nwholemount stain for axons.\n9. Figure 4, in chimeric animals what percentage of cells are Sox2-cherry NTR positive. Do the endogenous cells still\nrespond to the injury signal and proliferate or does NTR block the normal cell division.\\?\n10. Figure 5: telencephalon transplant experiment, it takes five months post-transplant, which only has a 16% success rate to\nbe able to then carry out an ablation, this again is not a useful experimental paradigm, especially since the morphology of\nthe brain is not normal after transplantation.\n11. Type of brain injury performed in Figure 5 neds to be described in the text.\n12. The authors then show that NTR2.0 works better and generate a nuroD6Cherry: NTR.0 line. However, in Figure 6 E, it\nstill appears to take 10 days of MTZ treatment before a significant decrease in cherry signal is observed. The text states that\na “massive depletion of cortical neurons at 5dpt” line 298-300). This statement needs to be supported by quantification of\nneuronal number in control versus treated samples by staining for NeuN and Sox2. If the ablation construct is specific to\nNeuroD in neurons, then the Sox2-positive glial cells in the dorsal pallium number should be unaffected in neuron-ablated\nanimals.\n13. Cortical layers in the axolotl brain have not been well defined. Control sections need to be shown where the markers\nused to define cortical layers are seen.\n14. Overall, the manuscript is lacking in quantification; no “N” is given for most experiments."}, "author_response": "Point to point response to the reviewers’ comments\nWe would like to express our sincere appreciation to the reviewers for their thorough\nevaluation of our work and their valuable feedback. All comments and suggestions\nprovided were highly constructive, and we have conducted additional experiments and\nanalyses in response. Our point-by-point responses are detailed below. With the newly\ngenerated experimental results and more comprehensive data analysis and\ninterpretation, we believe that we have fully addressed all the reviewers’ concerns and\nsubstantially improved our manuscript.\nDetailed point-by-point responses to specific reviewers' comments are provided\nbelow:\nReviewer #1:\nCell ablation tools are currently not available for studies using the axolotl, which\narguably is the best tetrapod model for regeneration research. The manuscript\nchronicles a series of experiments that establish efficacy in using NTR-NTR2.0/MTZ\ncell ablation to target neuronal cell types in the lateral line, spinal cord, and brain.\nAdditionally, the study reports development of a Cre-loxP-based reporter NTR2.0 line\nthat should allow targeted ablation of any axolotl Cre line. Although the manuscript is\nmethodologically oriented, several interesting biological findings are reported to\nestablish broader significance of the work, including Sox2+cell ablation that inhibited\nspinal cord and tail regeneration, and Neurod6+ cell ablation that provided an\nopportunity to detail neuronal patterning in the brain of a regeneration competent model.\nImpressive amount of work accomplished using the axolotl. I list some minor issues\nbelow.\nThank you for your positive feedback and constructive comments on our manuscript.\nWe have carefully revised the manuscript in accordance with your suggestions, and our\ndetailed responses are provided below.\nLine 40: I think \"chronologically\" is the correct word, and can you please describe more\nprecisely what is meant by layer organization.\nThank you for your valuable comment. We agree that \"chronologically\" is the correct\nword in this context, and we appreciate your suggestion to clarify \"layer organization\".\nWhat we described as \"layer organization\" refers to the laminar-like distribution of\nsequentially labeled regenerated neurons in the axolotl pallium. Specifically, through\nsequential BrdU/EdU labeling, we observed that regenerated neurons display a\nstereotypic laminar-like arrangement that carries chronological information: earlier-\nborn regenerated neurons (BrdU-labeled) are predominantly localized to the outer part\nof the pallium, while later-born ones (EdU-labeled) populate the inner part, forming an\noutside-in stratified pattern. Notably, this regenerative distribution mirrors the neuron\ngeneration pattern during embryonic development, indicating that neurogenesis and the\nformation of this laminar-like arrangement follow the same spatiotemporal sequence.\nWe have revised all relevant text to replace \"layer organization\" with \"laminar-like\ndistribution\" or \"zones\" and incorporated corresponding clarifications into the text to\nimprove accuracy.\nLine 41-42: Perhaps change the sentence to this: \", enabling tissue-specific ablation of\nspecific cell types when crossed to existing Cre lines.\nThank you. Your suggestion is definitely a better phrasing. We have updated the text as\nyou recommended (lines 44-45 in the revised manuscript).\nLine 54: Maybe change to \"knowledge gaps hinder\".\nLine 61: Maybe change to \"using genetic approaches\".\nLine 79: Italicize \"E. coli\".\nLines 139, 217, 221: remove comma.\nLine 140: delete \" that from the\".\nLine 149: delete \"live imaging\".\nLine 151: change \"the identified macrophages\" to IBA1+ cells.\nLines 171 and 180: I think \"compared\" is the correct word.\nLine 232: maybe change \"on\" to in.\nLine 234. Maybe end sentence with a period after telencephalon, then start next\nsentence with Sox2+.\nLine 238. Maybe end sentence with a period after 45 dpt, then start next sentence with\nCHERRY signals were undetectable, indicating ablation of Sox2+ EGCs, while\nfluorescence in controls remained unchanged.\nLine 247: remove comma after BrdU.\nLine 252: maybe change \"wounds healing via\" with evidence of.\nLine 257: maybe change \"neurons-derived\" to neuronal.\nLine 268: change \"on\" to of.\nLine 289: change sentence maybe to this: These results established NTR2.0/MTZ as\nsuperior method for neuronal ablation.\nLine 294: change \"originated\" to originating.\nLine 297: change \"on\" to of.\nLine 322: change to nearly completely lost.\nLine 373: change \"on\" to of.\nLine 399: change \"enable\" to enables.\nThank you for your thorough review and insightful suggestions. We have implemented\nall suggested revisions. These changes have markedly enhanced the clarity and\naccuracy of the manuscript.\nLine 88-89: Please rewrite this sentence: \"While axolotls exhibit remarkable brain\nregeneration after trauma, therefore it serves as an ideal model for brain regeneration\nresearch.\"\nThank you for your suggestion. We have rewritten as follows:\n\"Axolotls exhibit remarkable brain regeneration after trauma, making them an ideal\nmodel for brain regeneration research.\" (lines 90-91 in the revised manuscript)\nLines 89-91: Many studies predating single cell sequencing established a primary role\nfor EGCs in spinal regeneration and probably should be referenced.\nThank you. We have now incorporated references predating single-cell sequencing\nwork that established the essential role of EGCs in spinal cord and brain regeneration\n(references 24 and 70-73 in the revised manuscript).\nLines 127-129: Maybe rewrite these two sentences as they do not read well.\nThank you. We have revised these two sentences as follows:\n\"To develop targeted cell ablation in axolotls using the NTR system, we targeted EGCs,\nwhich has previously been established as the cellular source of spinal cord and brain\nregeneration in salamanders. However, their precise contribution to central nervous\nsystem (CNS) regeneration remains to be fully elucidated.\" (lines 113-116 in the revised\nmanuscript)\nLine 249: include a reference establish CTIP1 as a marker of new born neurons.\nThank you for pointing this out. “CTIP1” in previous Line 249 was a typo, which we\nhave now corrected to “CTIP2”.\nRegarding to your kind consideration, our original phrasing may be misleading. We\nintended to referring that the specific neuronal subtype marker CTIP2, co-labeling with\nBrdU, helps identify newly generated neurons of that particular subtype after injury. To\nclarify this point, we have revised the sentence as follows and included a reference\nindicating CTIP2 as specific neuronal subtype marker:\n\"Additionally, a substantial number of BrdU+ cells co-labeled with the neuronal subtype\nmarker CTIP2, confirming the generation of new neurons after traumatic injury.\" (lines\n250-252 in the revised manuscript)\nLine 262: It is debatable that relatively rapid depletion of cortical neurons by cell\nablation provides a model for protracted cell loss seen im many degenerative neuronal\ndisorders. Maybe clarify/better rationalize the manipulation as a model.\nThank you for raising this critical comment.\nWe agree that rapid depletion of cortical neurons in our model differs from the\nprogressive/gradual neuronal loss observed in most degenerative disorders. We have\nrevised the text to clarify the rationale for our neuron ablation model as follows:\n\"Neuronal loss is the key feature of many CNS disorder. Restoration of lost neurons\nremains a great challenge. Therefore, establishing a neuronal ablation model is crucial\nto study the neuronal regenerative capacity. To this end, we generated NeuroD6:\nCherry-NTR knock-in axolotls …\" (lines 261-264 in the revised manuscript)\nLine 300: please rewrite \" that comprised near solely\".\nThank you for your suggestion. We have rewritten the sentence as follows:\n\"… and a significant reduction in the thickness of the dorsal pallium, which was\ncomposed almost entirely of SOX2+ EGCs following neuronal ablation at 5 dpt.\" (lines\n279-280 in the revised manuscript)\nLine 313: please clarify what is meant by the developmental effect. Also, were\nbehavioral changes observed in adult axolotls as reported in larval axolotls.\nThank you for pointing this out. The term \"developmental effect\" is indeed confusing.\nWe have removed this term in our revised manuscript.\nIn the original text, the term \"developmental effect\" referred to ongoing CNS\ndevelopment might intertwine with regenerative processes rapid growth state in CNS\nat early stage, making it difficult to distinguish between \"regeneration after ablation\"\nand \"normal developmental progression\".\nTo address this concern, we conducted our experiments in adult axolotls with mature\nCNS that are no longer undergoing active development. This allowed us to eliminate\ndevelopmental confounders and focus specifically on the regenerative response to\nneuronal loss in adult animals. We have updated the sentence as follows:\n\"To investigate the spatial organization and subtypes of regenerated neurons after\nlarge-scale neuronal ablation in the dorsal pallium, and to avoid developmental\nprocesses confounding regenerative responses, we carried out neuronal ablation in\nadult (15 cm in size from snout to tail tip) NeuroD6:Cherry-NTR2.0 axolotls and\nanalyzed the neuronal regeneration.\" (lines 295-300 in the revised manuscript)\nRegarding behavioral observations in adult axolotls, we observed dysphagia-like\nphenotypes (swallowing dysfunction) consistent with those reported in larvae. However,\ndue to the opacity of adult tissues, direct visualization of food retention in the oral cavity\nwas not feasible. We therefore evaluated eating behavior and observed that Neurod6-\nablated adults took longer time to swallow the food, comparing to the control adults,\nconsistent with swallowing dysfunction documented in larva.\nLine 358-360. Please rewrite this sentence to achieve clarity.\nThank you for your suggestion. We have rewritten the sentence as follows:\n\"When combined with a specific Cre transgenic line, the floxed stop cassette is excised,\nresulting in Cherry-NTR2.0 expression under the control of the constitutive CAGGs\npromoter, thereby enabling targeted ablation of specific cells.\" (lines 343-345 in the\nrevised manuscript)\nLine 408: I don't think the results of the study enabled modeling of neurodegenerative\ndisease, maybe tone this statement down.\nThank you for this critical feedback. As we stated above, our objective is to establish a\nmodel for studying CNS regenerative capacity following large scale neuronal loss,\nwhich is a pathological feature observed in most degenerative disorders. Hence, our\noriginal statement may have overemphasized the direct relevance between our animal\nmodel and neurodegenerative disease modeling, particularly as our model did not\nrecapitulate the complexity of these disorders. We have revised the sentences as follows:\n\"(2) large-scale cortical neuron ablation, which induces defined neuronal loss\ncharacteristic of neurodegenerative diseases, sheds light on the robust neuronal\nregeneration capability of the adult axolotl CNS in a regenerative context, with\nimplications for understanding regenerative deficits in neurodegenerative conditions.\"\n(lines 394-397 in the revised manuscript)\nLines 438-449: Maybe good to cite again/comment on the more recent approach\nKID/KIS as this mediates cell-ablation more rapidly in zebrafish embryos than NTR2.0.\nThank you for highlighting the relevance of the Kid/Kis cell ablation system, which\nrepresents a valuable advancement in rapid and controllable cell elimination techniques.\nWe have incorporated a discussion of this method into our manuscript.\n\"NTR-mediated specific cell ablation exhibits temporal heterogeneity and is\nprogressive. This is primarily because the substrate needs to first diffuse into NTR-\nexpressing cells and accumulate to a threshold level to effectively trigger cell apoptosis,\nand this temporal course is closely associated with the NTR expression level and\nproliferative status of targeted cells. Notably, the recently developed Kid/Kis system\nrepresents a powerful alternative for rapid cell ablation, particularly in zebrafish\nembryos. This system utilizes bacterial toxin/anti-toxin proteins to mediate cell-\nautonomous elimination with significantly faster kinetics, thereby enabling\ninvestigations into the acute consequences of cell loss. Based on the principle of the\nKid system, it requires an inducible expression system to control cell ablation, such as\nthe widely used GAL4/UAS system in zebrafish and Drosophila, which has not yet been\nestablished in axolotls. However, the future integration of the Cre-LoxP system which\nhas already proven effective in axolotls with the Kid system could enable precise and\nrapid cell ablation, offering exciting possibilities for regeneration studies.\" (lines 438-\n451 in the revised manuscript)\nLines 455-458: Please rewrite this sentence to achieve clarity, and perhaps cite relevant\nresearch supporting the idea of low Cre conversion rate.\nThank you for your suggestion. We have revised this sentence and inserted relevant\ncitation.\n\"However, even when inducible Cre expression is driven by a strong promoter, Cre-\nmediated recombination of floxed alleles is generally difficult to reach 100% efficiency,\ntherefore, attenuating the complete ablation of the entire population of cells of interest.\"\n(lines 458-461 in the revised manuscript)\nLines 460-466: There are several small grammatical errors in this paragraph, for\nexample on line 464, maybe change \" targeted cells ablation are\" to \"cell ablation is\".\nThank you. We have carefully reviewed and revised accordingly.\nLines 482-484: How will the lines created in the study be made available for use by\nother researchers? Will they be donated to a stock center? This would seem important\nto increase axolotl research efforts community-wide.\nThank you for emphasizing the importance of sharing transgenic lines and resources\nwith other researchers in the field. We strongly agree with the reviewer on this point.\nAll transgenic axolotl lines generated in this study will be available to the axolotl\ncommunity. Following the reviewer’s suggestion, we will deposit relevant transgenic\nlines to the Ambystoma Genetic Stock Center (AGSC) at the University of Kentucky.\nWe have also included a statement on resource and data availability in the revised\nmanuscript.\nLines 546 and 563 and 574 and 631: What volume amounts were injected?\nThank you for pointing out this important detail. We have included the specific injection\nvolume information to the Materials and Methods (lines 548, 564, 565, 577, 644 and\n645 in the revised manuscript).\nLine 600: How exactly was the spinal cord removed? Can more detail be provided?\nThank you for reminding us to provide more details regarding the spinal cord excision\nand transplantation procedure. It is important for others to repeat this procedure. The\nmethodological description has been elaborated in the Materials and Methods in the\nrevised manuscript.\n\"We used donor Sox2:Cherry-NTR and host d/d axolotls (4 cm in body length) for spinal\ncord transplantation. All animals were deeply anesthetized prior to surgery. Both donor\nand host animals were amputated at the equivalent somite level (12–14th somite rostral\nto the cloaca) to expose the amputation plane. For the host, A lateral incision was made\nthrough the skin at 2 somites rostral to the amputation plane using fine surgical scissors,\nand the spinal cord was transected at this site. A 2-somite-length spinal cord segment\nwas then carefully extracted from the spinal canal through the amputation plane using\nsterile forceps, with gentle handling to preserve structural integrity. For the donor, A\n3-somite-length spinal cord segment at the equivalent anatomical site was removed\nusing the same extraction protocol: a lateral skin incision near the amputation plane,\ntransection of the spinal cord, and extraction of the target segment from the spinal canal\nvia the amputation plane. After extraction, the damaged end of the donor segment was\nslightly trimmed to refine the graft for transplantation. Immediately following host\nspinal cord removal, the donor segment (trimmed to match the 2-somite-length of the\nhost’s excised segment) was inserted into the host’s resulting spinal canal cavity. This\nensured precise anatomical alignment and direct contact between the transplanted graft\nand the host’s spinal cord stumps. Two months after spinal cord transplantation (at\n7cm in body length), the success of spinal cord transplantation was defined by normal\nmorphology in the regenerated tail segment, in which presented the donor-derived\nCHERRY positive spinal cord.” (lines 598-616 in the revised manuscript)\nReviewer #2:\nIn this manuscript, Fu et al develop the nitroreductase system for ablation of glial\ncells and neurons in the central nervous system in axolotls. This is a widely used\nsystem in zebrafish and mice, and could, in theory could be a very useful system to\nadapt in axolotls to ablate certain cell populations specifically. The manuscript is long,\nshowing a lot of things that did not work efficiently. Overall, the manuscript could be\nshortened and transformed into a more concise Methods paper. However, I have some\nmajor concerns about the ablation efficacy shown in the manuscript. Specific points:\nThank you for your constructive evaluation of our work and your recognition of the\nNTR-based ablation system's value in axolotls. We have carefully addressed your\nconcerns regarding manuscript structure and ablation efficacy.\nFollowing your suggestion, we have shortened the manuscript by moving several\nsections to the Supplementary Information, resulting in a more concise main text. This\nincludes the removal of the electroporation-based NTR data (previous Fig. 1) and the\ntransfer of NTR substrate optimization and NeuroD6:Cherry-NTR characterization to\nSupplementary results. These elements were retained in the supplement as they may aid\nreproducibility and prevent redundant efforts by other researchers.\nWe have updated the manuscript with sharpen narrative, focusing on: (1) establishing a\nvalidated NTR/NTR2.0 ablation system for axolotls; (2) revealing novel insights of\ncentral nervous system regeneration, including the essential role of EGCs and neuronal\nsubtype regeneration; (3) providing actionable NTR2.0-based genetic tools (knock-in\nlines, Cre-LoxP system) for the research community.\n1. The authors first test a ubiquitous promoter CAGG driving Cherry NTR and quantify\nfluorescence intensity in low mag images and show a decrease at 6 days post-treatment.\nThis does not prove that these cells types are actually ablated. I would remove these\npanel Figure 1a-e and the text.\nThank you for your critical comment. We apologize for this overstatement. We agree\nthat current data without sectioning and detailed characterization is insufficient to prove\nactual ablation of targeted cells. Following your suggestion, we have removed Figure\n1a-e and related text.\n2. The authors then generated a Sox2-Cherry-NTR line, these panels could be moved\nto Supplementary data.\nThank you for your suggestion. We have relocated the panels of Sox2:Cherry-NTR line\nto the Supplementary Information (Revised Supplementary Fig. 1).\n3. Figure 2 is confusing /unconvincing. Panel A shows that at 2 and 5 days post-\ntreatment with MTZ, there is lower fluorescence in the lateral line cells, but it is still\ndetected. But Panel C at 3dpt appears to show almost no Sox2+ cells after 3 days\ntreatment, however, the area of skin shown is much thinner than control, also like it also doesn’t have any lateral line\ncells here. This result needs quantification, whereby number of cells per neuromast that are expressing Sox2 are counted\nprior to giving MTZ, then the number of Sox2 positive cells counted 1dpt, 2dpt, etc. Number of animals used needs per\nexperimental condition needs to be stated in the Figure legend.\nThank you for your feedback and valuable suggestions. In response to your concerns,\nwe have conducted additional experiments and revised both the figure and manuscript\naccordingly. We provide our clarifications and the revisions as below.\nThe Cherry fluorescence observed in live images of Sox2:Cherry-NTR neuromasts\n(Revised Fig. 1a) does not represent intact SOX2+CHERRY+ cells, but rather the\nresidual CHERRY+ cellular debris from ablated SOX2+ cells that have been\nphagocytosed by macrophages but not yet fully degraded. This interpretation is\nsupported by co-staining of CHERRY and the macrophage marker IBA1, on both cross-\nsections (Revised Fig. 1g) and whole-mount immunofluorescence (Revised\nSupplementary Fig. 2f and Revised Supplementary Video 2).\nRegarding the apparent skin thinning following SOX2⁺ cell ablation, we attribute this\nto the collapse of the neuromast structure. The lateral line neuromast is a specialized,\nprotruding cup-shape structure in the skin (Revised Supplementary Video 1), containing\na core population of SOX2⁺ cells that are essential for maintaining its architectural\nintegrity. Ablation of these SOX2⁺ cells led to neuromast structural collapse (Revised\nSupplementary Video 2, Revised Supplementary Fig. 2c, d), which lead to reduces the\nthickness of the region similar to the surrounding skin.\nFollowing the reviewer’s suggestion, we performed whole-mount imaging and\nquantified the number of SOX2⁺ cells in neuromasts at 0, 1, 2, 3, 4, and 5 days post-\nMTZ treatment (Revised Supplementary Fig. 2e). The results demonstrated a\nprogressive reduction of neuromast SOX2⁺ cells, with approximately 22% loss at 1 dpt,\n67% at 2 dpt, and over 90% by 3 dpt, compared to the baseline at 0 dpt. Additionally,\nthe number of SOX2+ neuromast cells started to increase from 4 days post-MTZ\ntreatment relative to controls. It presumably due to the initiation of neuromast\nregeneration, as our observed in live images (Revised Supplementary Fig. 4a).\nMeanwhile, with these recommended analyses, we identified a subset of weakly\nSOX2+ cells adjacent to the cup-shaped structure that appeared refractory to ablation,\nin both sectional and whole-mount staining. Therefore, we have replaced a more\nrepresentative image in Revised Fig. 1c.\nWe have included both the figures and relevant text in the revised manuscript. (Revised\nSupplementary Fig. 2, lines 129-147 in the revised manuscript)\n4. Figure 3 the authors determine that NFP is an effective drug for ablation of Sox2 glial\ncells in the spinal cord. However it is very concerning that it takes 10 days of treatment\nfor 51 of the cells to be ablated. When 50% of cells are ablated do the surrounding glial\ncells begin to proliferate to compensate for the dying cells?\nThank you for your insightful question. This is a great point. We have performed new\nexperiments to examine the proliferation in spinal cord after 10 days post-NFP\ntreatment and integrated new data into the revised manuscript.\nFor the first concern from the reviewer, “it takes 10 days of treatment for 51 of the cells\nto be ablated”. We treated axolotls with a single 24-hour pulse of the prodrug (MTZ or\nNFP) to ablate targeted cells (EGCs). Comparing MTZ with NFP, MTZ failed to ablate\nthe spinal cord EGCs; in contrast, administration of NFP resulted in loss of EGCs.\nAlthough only 49% of targeted cells was ablated by 10 days post-NFP treatment, it\nreached 94% at 20 days post-NFP treatment (Revised Fig. 2e). We admit this is a rather\nslow process. However, it has been reported that the performance of NTR/MTZ-\nmediated cell ablation varies in zebrafish, took between 1-3 days depending on the\ntargeted cell types(Curado et al., 2008; Mathias et al., 2014). The gradual depletion of\nEGCs with NFP might reflect the deep location of EGCs in tissues and level of NTR\nexpression. However, it might due to the overall insufficiency of NTR system compared\nto NTR2.0. Perhaps creation of a Sox2:Cherry-NTR2.0 line and combine with MTZ\ntreatment could achieve a faster spinal cord EGC ablation. To be precise, we have\nrevised the term from efficient to effective when describing NFP.\nFollowing the reviewer’s suggestion, we performed EdU incorporation assays to assess\nthe proliferative state of remaining SOX2+ cells after EGC ablation. Specifically, we\nadministered a single pulse of EdU 6 hours prior to sample harvest, followed by\nimmunofluorescence analysis. The results showed that about 10% of SOX2+ cells were\nBrdU+ in the spinal cord of DMSO-treated control animals (Revised Supplementary\nFig. 5a, c). In contrast, no BrdU+ SOX2+ cells were detected in the spinal cord at 10\ndays post-NFP treatment (Revised Supplementary Fig. 5b, c). These data indicated that\nthe residual SOX2+ EGCs did not actively proliferate to compensate for dying cells.\nThe enzyme NTR catalyze the substrates (e.g., MTZ or NFP) into toxic product to\ncrosslink the genomic DNA. The SOX2+ EGCs observed in the spinal cord at 10 days\npost-NFP treatment, although still alive, they failed to proliferate due to genomic DNA\ncrosslinking. It ultimately leads to ablation of these cells as observed at 20 days post-\nNFP treatment. We have added this information to revised manuscript as follows:\n“Immunostaining on tail and brain sections revealed structural disruption and dramatic\nreduction of SOX2+ EGCs after NFP treatment (Fig. 2a, b, j), compared to MTZ-treated\n(Fig. 2c, k) or DMSO-treated controls (Fig. 2d, l). The number of the spinal cord EGCs\ndecreased to 51.3% at 10 dpt (Fig. 2e), with no proliferative compensation in the\nresidual SOX2⁺ population (Supplementary Fig. 5a-c), and a further reduction to 5.5%\nby 20 dpt (Fig. 2e).” (lines 162-167 in the revised manuscript)\n5. The fact that the authors see 50% decline in spinal cord neurons at 20dpt is not\nsurprising since these animals are still growing and the glial cells give rise to new\nneurons and sox2 is a neural progenitor marker.\nThank you. We fully agree with your interpretation. Since the axolotls used in our\nablation experiments are larvae and continuously growing, Sox2+ EGCs (neural\nprogenitors) sustain ongoing neurogenesis, thus, their ablation logically leads to the\nobserved 50% reduction in spinal cord neurons at 20 dpt.\n6. In Figure 4 If they administer NTR for 20 days to ablate the Sox2 positive cells in\nthe spinal cord, then the animals die, this overall suggests that bathing the animals in\nthe drug for 20 days maybe the cause of death and not the ablation of the cells. In the\ntest of nonspecific toxicity of the drug they only tested to 10 days post treatment, a\ncontrol for 20 dpt needs to be done. Have you tried injecting the drug into the lumen of\nthe spinal cord instead of bathing the animals?\nThank you for your consideration and concern. In our study, for Sox2:Cherry-NTR\naxolotls, we treated axolotls with a single 24-hour pulse of the prodrug (MTZ or NFP)\nto ablate targeted cells (EGCs); for spinal cord-transplanted d/d animals, we treated\nthem with four 24-hour pulses, each pulse separated by a 24-hour interval. Both\nregimens were optimized based on preliminary tests, which confirmed that they are safe\nfor d/d axolotls and effective for cell ablation for Sox2:Cherry-NTR tissue, without\ninducing mortality or signs of toxicity (e.g., normal food intake and movement).\nMoreover, following the reviewer’s suggestion, we found that prolonged NFP treatment\nto d/d axolotls did not cause lethality (100% survival, Revised Supplementary Fig. 3c)\nand no sign of toxicity in d/d axolotls up to 20 dpt.\nIn summary, our results demonstrated that the NTR substrate does not cause mortality\nor obvious toxicity in d/d axolotls at our regimen. The lethality observed in\nSox2:Cherry-NTR axolotls is presumably linked to the specific ablation of Sox2+ cells,\nunderscoring their essential role in tissue homeostasis and regeneration.\nFurthermore, we have tried injecting the drug into the lumen of the spinal cord of\nSox2:Cherry-NTR axolotls. In pilot trials, intraluminal injections of MTZ at 40 mM\nproduced no detectable ablation (Response Fig. 1a, b). Considering the axolotl spinal\ncanal is extremely narrow, restricting injected volume to the microliter range (< 5 µL).\nThis small volume may disperse rapidly into cerebrospinal fluid and circulation,\nyielding only transient local drug exposure that was insufficient to trigger effective\nNTR-mediated ablation of Sox2+ cells. Moreover, attempts to increase MTZ stock\nconcentration for injection were not feasible. MTZ exhibits limited solubility, and\nhigher concentrations lead to immediate crystal precipitation during injection, clogging\nneedles and causing inconsistent delivery. Given these limitations, intraluminal\ninjection of prodrug is not feasible for cell ablation in the axolotl spinal cord.\nResponse Fig. 1: Lumen injection of MTZ leads to no ablation in Sox2:Cherry-\nNTR axolotls.\na Representative immunofluorescence images of spinal cord cross-sections from lumen\ninjected Sox2:Cherry-NTR axolotls at 10 days post-treatment (dpt), stained with SOX2\nand combined with DAPI. b Quantification of SOX2+ cells in lumen injected\nSox2:Cherry-NTR axolotls. Five serial sections from equivalent somite positions were\nanalyzed (n = 3). Each dot represents an individual animal’ mean value. Data were\nanalyzed by unpair two-tailed Student’s t-test and represented as mean ± SEM. ns\nindicates P > 0.05. Scale bars: 50 μm in a.\n7. In Figure 4 the authors embark on a complicated unnecessary experiment to show\nthat glial cells are necessary for spinal cord regeneration. The authors create chimeric\nanimals by spinal cord transplantation. The better experiment would be to simply treat\nfor 5 days with NTR, amputate the tail, continue treatment for 5days, whereby at least\n50% of the cells should be ablated and measure the length of the regenerate overtime\nversus the control animals.\nThank you for your comment. As stated in our responses to Reviewer 2 (question #4\nand #6), a single 24-hour pulse of NFP treatment effectively ablated Sox2+ cells in\nSox2:Cherry-NTR axolotls (Revised Fig. 2). As Sox2 is expressed throughout the\ncentral nervous system and several other organs in axolotls, its ablation affected\nsystems beyond the CNS. Consequently, although all treated axolotls survived until 20\ndpt (the peak of EGC ablation), they subsequently exhibited abnormal phenotypes,\nincluding a belly-up posture, body curvature, and impaired mobility, accompanied by a\ncessation of feeding. Notably, from day 20 onward, gradual mortality was observed in\na subset of these treated animals.\nGiven the lethality and unhealthy state of the NFP-treated axolotls, it is difficult to\nconclude that observed regeneration defects are caused solely by the loss of SOX2+\nEGCs, as potential systemic effects cannot be ruled out. Hence, spinal cord\ntransplantation is essential for studying the function of EGCs post-ablation.\nIn response to your suggestion, we performed a direct treatment experiment in\nSox2:Cherry-NTR axolotls following the proposed timeline. However, this approach\nencountered critical issues related to animal survival and treatment specificity. When\nSox2:Cherry-NTR axolotls received a continuous treatment with 7.5 μM or 5 μM NFP,\nall animals died by 4 or 6 days post-treatment, respectively, no subsequent regeneration\nanalysis can be proceeded (Response Fig. 2a). When transgenics received a continuous\ntreatment with 2.5 μM NFP, animals showed a noticeable delay of regeneration but still\nretained the ability of spinal cord regeneration (Response Fig. 2a, b, c). We suspect this\ndelay may reflect insufficient Sox2+ cell ablation. Therefore, we underscore the\nlimitations of directly treating Sox2:Cherry-NTR animals and reaffirm the necessity of\nusing transplantation experiments to definitively investigate the role of Sox2+ EGCs in\nspinal cord regeneration after EGCs ablation.\nResponse Fig. 2: Tail regeneration followed by EGC ablation in Sox2:Cherry-\nNTR animals.\na The experimental timeline of NFP treatment in Sox2:Cherry-NTR axolotls. Animals\ntreated continuously with 7.5 μM or 5 μM NFP were succumbed at 4 and 6 dpt,\nrespectively, whereas those treated with 2.5 μM NFP survived through the observation\nperiod. Red crosses indicate time points of complete mortality. b Representative time-\nseries images of tail regeneration in Sox2:Cherry-NTR axolotls following continuous\n2.5 μM NFP or DMSO treatment. NFP-treated animals (top) exhibited delayed spinal\ncord regeneration compared to controls (bottom). c Quantification of the regenerated\nspinal cord length in Sox2:Cherry-NTR axolotls after continuous 2.5 μM NFP or DMSO\ntreatment. Length was measured from amputation plane to CHERRY+ spinal cord tip at\n0, 5, 10, 15, 30 days post amputation (n = 3 in each group). Each dot represents an\nindividual animal. Dashed lines indicate the amputation plane, and arrows indicate the\nCHERRY+ spinal cord. Data were analyzed by unpair two-tailed Student’s t-test and\nrepresented as mean ± SEM. ***p<0.001. Scale bars: 2 mm.\n8. Line 219, referring to Figure 4, \" neuronal loss likely contributed to this phenotype\".\nThe authors need to quantify neuronal regeneration in control versus ablated animals\nby carrying out NeuN staining on sections and quantifying and doing a wholemount\nstain for axons.\nThank you for this valuable suggestion. In response, we have performed quantitative\nanalyses on neurons on sections and examined axons with wholemount stain.\nFirst, we carried out immunostaining and quantification on spinal cord sections from\nspinal cord transplanted animals at 20 dpt to analyze regenerated neuron and axon.\nWhile SOX2+ EGCs were nearly absent following NFP treatment (Revised Fig. 3h- j),\nwe observed an approximately 35% decrease in NeuN+ neurons and an approximately\n23% reduction in TUJ1+ axonal fluorescence intensity compared to controls (Revised\nFig. 3k–p).\nSecond, to assess axons distribution in the regenerated tails, we performed IDISCO\nwholemount staining as your suggestion. In NFP-treated chimeras, both SOX2+ cells\nand TUJ1+ axons were sparse in the regenerated tail tip region at 10 dpa (Revised\nSupplementary Fig. 6a, b and Revised Supplementary Video 3).\nIn contrast, control\nanimals exhibited abundant SOX2⁺ cells in the spinal cord and TUJ1⁺ axons extending\nto the tail tip (Revised Supplementary Fig. 6a, b and Revised Supplementary Video 4).\nOur results link the loss of EGCs, along with impaired neurons and axons, resulted in\nthe observed regeneration deficit. We have incorporated these findings into revised\nfigures (Revised Fig. 3j-m, Revised Supplementary Fig. 6a, b, and Revised\nsupplementary Video 3, 4). Additionally, we have updated the relevant text as follows:\n\"Additionally, we observed a 35% reduction in NeuN+ neurons and a 23% reduction in\nTUJ1+ fluorescence intensity at 10 dpa compared with controls (Fig. 3k-p). Whole-\nmount immunostaining of regenerating tails from NFP-treated chimeras further showed\nboth SOX2+ cells and TUJ1+ axons were sparse in the regenerated tail tip region, while\ncontrol animals exhibited abundant SOX2⁺ cells in the spinal cord and TUJ1⁺ axons\nextending to the tail tip (Supplementary Figure 6a, b and Supplementary Video 3, 4).\nThese findings collectively indicate that neuronal loss likely contributed to regeneration\ndeficits in both the spinal cord and tail.\" (lines 208-215 in the revised manuscript)\n9. Figure 4, in chimeric animals what percentage of cells are Sox2-cherry NTR positive.\nDo the endogenous cells still respond to the injury signal and proliferate or does NTR\nblock the normal cell division.\\?\nThank you for these critical questions. Following your question, we have performed\nadditional experiment and quantification to analyze cell composition and proliferative\ncapacity in chimeric animals.\nTo assess host EGCs contribution to transplanted spinal cord, we examined cross-\nsections from three spinal cord transplanted chimeras prior to NFP treatment. Donor-\nderived CHERRY+ cells accounted for 100% of SOX2+ cells in the transplanted and\nnewly regenerated spinal cord (Revised Fig. 3c, d), indicating no contribution from\nhost-derived SOX2+ cells (CHERRY-) to spinal cord transplants. Following NFP\ntreatment, approximately 93% of SOX2+ EGCs were ablated by 20 dpt (Revised Fig.\n3h-j). The remaining 7% SOX2+ cells were BrdU-negative (Revised Fig. 3h-j),\nindicating they failed to proliferate to replenish the ablated population (see also\nquestion #4 from the reviewer 2).\nTo examine the injury response of host (endogenous) EGCs, we quantified EGCs\nproliferation in the host spinal cord, near the interface of graft–host fusion (Revised\nSupplementary Fig. 6c, area indicating by A). There is a significantly increased\nproliferation in NFP-treated animals (~32.7% BrdU+SOX2+) compared to controls\n(~16.5%) (Revised Supplementary Fig. 6c–e). This localized proliferative response\noccurred >5 mm from the amputation site, beyond the typical range identified in\nprevious study, showing injury-induced activation is spatially restricted to ~500 μm to\nthe wound(Rodrigo Albors et al., 2015). It suggested that (1) loss of donor EGCs after\nNFP treatment did not affect the endogenous host EGC behavior; (2) massive loss of\ndonor EGCs induced the proliferative response of adjacent endogenous EGCs ablation\nrather than amputation.\nWe have added this information to the revised manuscript as follows:\n“We further assessed the proliferation of host EGCs adjacent to the graft-host interface\nfollowing the ablation of donor EGCs. Compared to controls (~16.5% BrdU+ in SOX2+\ncells), the host EGCs exhibited higher proliferating rate (~32.7%) in NFP-treated\nchimeras (Supplementary Fig. 6c-e). This localized proliferative response (>5 mm from\namputation plane) is beyond the typical range of injury-induced activation (~500 μm),\ntherefore we considered this host EGCs proliferation induced by massive loss of donor\nEGCs rather than amputation.” (lines 217-223 in the revised manuscript)\n10. Figure 5: telencephalon transplant experiment, it takes five months post-transplant,\nwhich only has a 16% success rate to be able to then carry out an ablation, this again is\nnot a useful experimental paradigm, especially since the morphology of the brain is not\nnormal after transplantation.\nThank you for this critical comment. We agree that the telencephalon transplantation\nexperiment is complicated, time-consuming, with some technical limitations, which we\nare actively working to improve.\nAs noted in our response to Reviewer #2 (Question #7), we believe telencephalon\ntransplantation remains essential for studying EGC function after ablation. Through this\napproach, we were able to demonstrate that EGC loss leads to complete abolition of\nnewborn EGCs, impairs injury-induced EGC proliferation and the generation of\nnewborn neurons (Revised Fig. 4j, i). It suggested that EGC is the sole source of neuron\nregeneration in axolotl, no other source to transdifferentiate to produce neuron in the\nabsence of EGCs.\nFurthermore, we would like to clarify that the 16% success rate reflects our strict\ncriterion in evaluating chimeras, requiring both graft survival and normal morphology\n(Response Fig. 3). The rate of chimeras maintaining partial CHERRY+ grafts is\nsubstantially higher (around 50%).\nDespite current limitations, we are actively refining this technique and believe it holds\nmore potential. Beyond the current study, combining with NTR/NTR2.0 and other\ngenetic tools, this approach could enable future studies in areas such as investigation of\nthe memory by brain tissue transplantation, offering new insights into neuroscience.\nResponse Fig. 3: Variable pattern in telencephalon transplants of chimeric\nSox2:Cherry-NTR axolotls.\nRepresentative images of variable telencephalic transplantation pattern (pattern 1-5) in\nchimeric Sox2:Cherry-NTR axolotls (from donor Sox2:Cherry-NTR into host d/d\naxolotl) at 5 months post-transplantation. Images are presented in dark field (top) and\ncorresponding CHERRY fluorescence (bottom). Total number of transplants: n = 36.\nThe number of transplants exhibiting consistent pattern is indicated below each\nrepresentative examples. Scale bars: 2 mm.\n11. Type of brain injury performed in Figure 5 neds to be described in the text.\nThank you for pointing this out. We have now included a detailed description of the\nbrain injury procedure in the Materials and Methods, as follows:\n\"Axolotl brain injury was conducted as previously described. Briefly, deeply\nanesthetized animals were made a rectangular cranial skin/skull flap over the left\ntelencephalon, using a scalpel and spring scissors and reflected to expose the\ntelencephalon. For the injury, a 0.5 mm × 0.5 mm square of tissue was excised from the\ndorsal region of the left telencephalon. To standardize the lesion site across animals,\nthe excision was precisely localized between the choroid plexus and olfactory bulb of\nthe left telencephalon. After injury, the cranial skin/skull flaps were repositioned\nwithout sutures.\" (lines 634-640 in the revised manuscript)\n12. The authors then show that NTR2.0 works better and generate a nuroD6Cherry:\nNTR.0 line. However, in Figure 6 E, it still appears to take 10 days of MTZ treatment\nbefore a significant decrease in cherry signal is observed. The text states that a \"massive\ndepletion of cortical neurons at 5dpt\" line 298-300). This statement needs to be\nsupported by quantification of neuronal number in control versus treated samples by\nstaining for NeuN and Sox2. If the ablation construct is specific to NeuroD in neurons,\nthen the Sox2-positive glial cells in the dorsal pallium number should be unaffected in\nneuron-ablated animals.\nThank you for these valuable suggestions. We have elucidated the identified CHERRY\nsignal composition following neuronal depletion and performed the recommended\nanalyses to evaluate the specificity of the ablation system.\nFirstly, the apparent slower CHERRY signal reduction in live imaging (Revised Fig. 5e)\nlikely reflects fluorescence from non-degraded CHERRY+ cellular debris engulfed in\nmicroglia, confirmed by co-staining with IBA1 at 5dpt (Revised Fig. 5j). This\nobservation is consistent with the phenotypes previously observed following lateral\nneuromast ablation.\nFollowing your suggestion, we next quantified neurons and EGCs in dorsal pallium of\nNeuroD6:Cherry-NTR2.0 animals. Since not all NeuN+ neurons are CHERRY+\n(Revised Fig. 5g), we specifically quantified CHERRY+NeuN+ cells. Our results\nrevealed a ~93% reduction of CHERRY+NeuN+ cells in MTZ-treated animals at 5 dpt\n(Revised Fig. 5h), confirming massive neuronal loss. In contrast, the quantification of\nSOX2+ EGCs showed no significant difference between controls and MTZ-treated\nanimals (Revised Fig. 5g, i), confirming neuronal‐specific ablation.\nThese results are consistent with your expectations and support the lineage relationship:\nwhereas EGCs ablation impairs neuronal production, neuronal ablation does not deplete\nEGCs but will promote EGC proliferation and differentiation upon neuronal ablation\n(Revised Supplementary Fig. 10m-o).\n13. Cortical layers in the axolotl brain have not been well defined. Control sections\nneed to be shown where the markers used to define cortical layers are seen.\nThank you. We apologize for the imprecise description. As noted in our response to\nReviewer #1 (Question #1), we previously described \"layer organization\" refers to the\nlaminar-like distribution of sequentially labeled regenerated neurons in the axolotl\npallium. We have revised all relevant text to replace \"layer organization\" with \"laminar-\nlike distribution\" or \"zones\".\nWe fully agree that cortical layers in the axolotl brain are not well defined. Currently\nno specific molecular markers distinguishing these layers have been established in the\nliterature or in our study.\n14. Overall, the manuscript is lacking in quantification; no \"N\" is given for most\nexperiments.\nThank you for your valuable feedback. We have thoroughly addressed the sample sizes\nand quantification throughout the manuscript.\nFor all quantitative analyses, we have included explicitly state the sample size (N,\nbiological replicates) in the figure legends, Methods and Material, and Source Data\ndocument, together with appropriate statistical tests.\nReferences\nCurado, S., Stainier, D. Y., & Anderson, R. M. (2008). Nitroreductase-mediated\ncell/tissue ablation in zebrafish: a spatially and temporally controlled ablation method\nwith applications in developmental and regeneration studies. Nature Protocols, 3(6),\n948-954.\nMathias, J. R., Zhang, Z., Saxena, M. T., & Mumm, J. S. (2014). Enhanced cell-specific\nablation in zebrafish using a triple mutant of Escherichia coli nitroreductase. Zebrafish,\n11(2), 85-97.\nRodrigo Albors, A., Tazaki, A., Rost, F., Nowoshilow, S., Chara, O., & Tanaka, E. M.\n(2015). Planar cell polarity-mediated induction of neural stem cell expansion during\naxolotl spinal cord regeneration. Elife, 4, e10230."}, {"round": 2, "reviewer_comments": {"reviewer_1": "I am satisfied with the authors responses to my concerns and changes made to improve the manuscript.", "reviewer_2": "Overall the authors ahve greatly improved the manuscript, both in the data they show and in the text.\nOne small point is that I dont beleive the title accurately describes the data they show. The data does not show \" the pivotal\nrole of ependymoglial in the entire cortex regeneration in axolotls\".\nThe entire cortex was never fully ablated in these experiments, the title should be modified to more accurately reflect the\ndata in the manuscript.\nAfter that, then the manuscript woudl be suitable for publication."}, "author_response": ""}]

Decision letter