Cellular response to injury in regenerative and non-regenerative stages
The cellular organization of the spinal cord CC in Xenopus laevis changes between regenerative and non-regenerative stages [48]. To determine the cellular response to spinal cord injury between regenerative (R-stages, NF stage 50) and non-regenerative (NR-stages, NF stage 66) stages, we performed a detailed cellular analysis.
The spinal cord of R-stage animals was injured by full transection as described previously [40] (Fig. 1A), and tissues were analyzed by light and electron microscopy at different days post transection (dpt). At 2 dpt (Fig. 1B,D), a complete sealing of the rostral stump was observed (Fig. 1B, arrowheads in Fig. 1D). The cells lining the CC close to the injury site were not affected by the lesion. To identify ultrastructural changes in CC cells after SCI, we analyzed ultrathin sections. Cells lining the CC, characterized in the control as type I, II or III [48], lack junction complexes compared to controls (Fig. 1E, arrowheads), contain swelled mitochondria in their apical pole (Fig. 1E, arrow), and frequent centriolar satellites were found (see supplementary material, Fig. S1A, arrowheads). As expected, we identified abundant cells showing mitotic figures indicating cell division [46–47], almost half of the cellular clusters undergoing cell division have no contact with the central canal lumen (Fig. 1F), while the other half are in direct contact with it (Fig. 1G). Although in a lower proportion, cell division in the CC has been also observed in uninjured animals [47–48]. Conspicuous among the cells lining the CC was the presence of donut- and phone-like shaped mitochondria, phenome not observed in control animals (see supplementary material, Fig. S1B, arrowheads). Of note, we found cells in the CC that were being extruded toward the lumen (Fig. 1B,H arrowheads). Regarding infiltration in the injury site we found very few red blood cells (data not shown), abundant macrophages (Fig. 1C, arrowheads) with an electron dense cytoplasm (Fig. 1I), and a high number of ribosomes (data not shown) that are phagocytosing cellular debris (Fig. 1I), and some neutrophils (see supplementary material, Fig. S1C) that were not observed in controls.
At 6 dpt cells accumulate in the lumen of the central canal (Fig. 1J,K). These cells showed a higher nucleus/cytoplasm ratio similar to the cells lining the CC, a lax chromatin, a scarce cytoplasm with few organelles, and the absence of cellular junctions between them or with cells of the spinal cord (Fig. 1K, arrows). Interestingly, some cellular expansions within the lumen exhibited a high density of light vesicles, and few small dense core vesicles (Fig. 1L, white arrowheads) in near proximity to structures reminiscent of postsynaptic densities (Fig. 1L, black arrowheads). At this time, clusters of 20–24 cells forming rosette like structures were found in the ablation gap (Fig. 1J, M). These cells are very similar to those found in the uninjured central canal; resembling type III cells previously described [48]. These cells have a characteristic neuroepithelium organization; with a basal lamina containing collagen (see supplementary material, Fig. S1D), a high nucleus/cytoplasm ratio, apical mitochondria, abundant apical interdigitations with adherent cell junctions (see supplementary material, Fig. S1E), a high content of intermediate filaments (see supplementary material, Fig. S1F), and the presence of a cilium (data not shown).
At 10 dpt some continuity between the rostral and caudal stumps was observed (Fig. 1N). Surprisingly, abundant bundles of axons were found inside the lumen of the caudal central canal (Fig. 1N, O). These bundles were mainly composed by unmyelinated axons (Fig. 1P, orange) that were in close contact with synaptic vesicles (Fig. 1P, arrowheads), but also some myelinated axons were detected (Fig. 1R, arrowhead). The axon bundles were surrounded by cells with a fusiform nucleus that have a lax chromatin (Fig. 1O, arrowheads), and desmosome-like junctions were found between those cells (Fig. 1Q). These cells in close contact with axons have a condensed chromatin resembling a neuronal morphology (Fig. 1S). In line with the observation at 6 dpt, rosettes and some cells are still present in the ablation gap (data not shown), and the central canal, respectively.
Finally, as described before [47] at 20 dpt we observed an almost complete reconstruction of the spinal cord (Fig. 1T). However, the continuity of the central canal was not perfect (Fig. 1T, arrowhead). Importantly, the cells lining the central canal recover its normal organization as a pseudo-stratified epithelium (Fig. 1U), with apical mitochondria, microvilli (Fig. 1V), and desmosome cell junctions (Fig. 1W). Few cells remained inside the lumen of the central canal (Fig. 1U, red line).
A similar analysis was carried out in NR-stage animals. For this the spinal cord of animals at stage 66 was transected as described [40] (Fig. 2A), and the CC was analyzed at different times after injury. Contrary to the response observed in the R-stage, at 2 dpt the cells lining the CC were severely damaged, with the presence of many empty spaces between cells, leading to a strong disorganization and loss of the pseudo-stratified epithelial organization (Fig. 2B,C, arrowheads). An important loss of intracellular organelles is observed in the remaining ependymal cells (data not shown). In addition, we observed a massive invasion of the ablation gap with blood cells, (Fig. 2B,D; red shadow), and the deposition of extracellular matrix (ECM) components (Fig. 2D, arrowheads). A massive disorganization of the central canal, is still observed at 6 dpt, together with a sustained increase in the extracellular spaces between cells, and the presence of vacuolated cells (Fig. 2E,F; see arrowheads). In addition, a massive presence of red blood cells, macrophages (Fig. 2E,G), and microglia (data not shown) was still detected in the injury site.
At 10 dpt, the CC cells have recovered some epithelial organization (Fig. 2H,I), and have an abundant number of mitochondria in the apical surface (Fig. 2I, J) which resemble the lateral ependymal cells described before [48]. Unlike the response at the R-stage, froglets at the NR-stage are characterized by the absence of proliferation and rosette-like structures. Ten days after injury the borders of the rostral and caudal stump were surrounded by glial processes (Fig. 2K, white arrowheads), containing abundant intermediate filaments (Fig. 2L, white arrowheads). Finally, at 20 dpt the presence of red blood cells, and immune cells in the injury site had decreased (Fig. 2M), and the ablation gap is mainly filled by fibroblast-like cells, characterized by long nuclei (Fig. 2N, white arrowhead), a very dilated rough endoplasmic reticulum (Fig. 2O, white arrowheads), and an ECM containing abundant collagen fibers (Fig. 2P). These morphological features correlate with the complete lack of swimming capacities at 20 dpt in NR-stage [46–47].
In summary, R-stages revealed a dynamic regenerative process characterized by a fast response of cells lining the central canal to rapidly seal the injured stumps, activating a proliferative response followed by formation of rosette-like structures in the ablation gap. In addition, cells are extruded into the lumen of the central canal, which is also filled with axons and synaptic vesicles. Regenerated spinal cords, although not always with a perfect morphology, indicates an efficient resolution of the regeneration at 20 dpt. On the contrary, in the NR-stage, cells lining the central canal are deeply affected after injury, and instead, red blood cells and macrophages populate the injury site. After several days, cells lining the canal recover their ultrastructure, but the injury site is filled with glial cells processes, fibroblast and collagen fibers, reminiscent of the glial scar described in mammals, and a proper recovery of the spinal cord was not observed.
Analysis of the presence of glial scar markers in response to spinal cord injury
One of the hallmarks of the cellular response to SCI in mammals is the formation of a glial scar composed of a fibrotic scar, and an astroglial scar border [6]. This scar is composed of different cell types including astrocytes, microglia, pericytes, and inflammatory and meningeal cells, together with ECM components such as fibronectin, CSPGs, and collagen among others [17–18]. To evaluate the formation of a glial scar in R- and NR-stages in response to injury we evaluated the presence of some of these markers at different days after injury.
First, we studied the expression of vimentin, an intermediate filament that is a marker of glial cells. In R-stage, vimentin was found in radial filaments located at the dorsal domain in the uninjured spinal cord (Fig. 3A, white arrowheads), as previously described (Edwards-Faret et al., 2018). Two days after injury, there was an increase in the number of cells expressing vimentin especially in the ablation gap and the regions close to the injury site (Fig. 3B, white arrowheads). Although a decrease in cells expressing vimentin was observed at 6 dpt, they are still higher than those detected in uninjured animals (Fig. 3C). For a more quantitative analysis, the region of the spinal cord surrounding the injury site was isolated and homogenized for western blot analysis. In this analysis, no change of vimentin levels was observed at 6, 10 and 20 dpt (Fig. 3D, see supplementary material, Fig. S2A). In NR-stage froglets, vimentin was only expressed in blood vessels in uninjured froglets (Fig. 3E-E’). At 10 dpt, vimentin was detected in cells with radial processes at the edges of the lesion (Fig. 3F-F’, withe arrowheads) and later, at 20 dpt, many of the vimentin positive cells with radial processes were now present in the ablation gap, most probably representing glial cells forming a glial scar (Fig. 3G-G’). The observed rise in vimentin correlates with the increase in radial processes previously detected by EM analysis, in which we observed processes with intermediate filaments at the edge of the lesion (Fig. 2K-L). This increase in the levels of vimentin at later days was confirmed by western blot analysis (Fig. 3H, see supplementary material, Fig. S2B).
The presence of the ECM components typical of the glial scar in mammals was evaluated. Fibronectin is expressed in the meninges in uninjured conditions in R-stage (Fig. 3I). However, at 6 dpt a clear increase in fibronectin was detected in cells that seal the rostral and caudal stump, most probably corresponding to meningeal cells, and was also found in more cells in the injury site (Fig. 3J). This increase is transient, and the levels of fibronectin return to almost normal at 10 dpt, being only expressed in the meningeal layer of the spinal cord (Fig. 3K). Fibronectic was almost not detected in froglets (Fig. 3L), and a similar response but more delayed, was observed. At 10 dpt, there is an increase in fibronectin deposition in the lesion site in the rostral and caudal stumps (Fig. 3M), and the levels are normal around 30 dpt (Fig. 3N). Similarly, we performed analysis of CSPGs, in NR-stages. Uninjured froglets showed expression of CSPGs only in blood vessels and vertebrae (Fig. 3O), and after injury a clear increase in the lesion site was observed, and was still present at 40 dpt (Fig. 3P-P´). CSPGs were not detected in the spinal cord before or after injury in R-stage animals (data not shown). For further analysis of collagen deposition, spinal cord sections were stained with Acid Fuchsin Orange G (AFOG), which labels collagen in blue, cells in orange and fibrin in red. Collagen was expressed in the meningeal layer in uninjured R-stage and froglets (see arrowheads, Fig. 3Q and T-T’). However, at 6 dpt the levels of collagen increased in the lesion site in R-stage (Fig. 3R), and at 10 dpt in froglets (Fig. 3U-U’). Interestingly, in R-stages the levels of collagen decreased, at 10 dpt (Fig. 3S), but high levels of collagen were still present in the lesion site in froglets at 20 dpt (Fig. 3V-V’).
For an unbiased comparison of the expression of glial markers in response to injury in R- and NR-stages we explored a data set from a high-throughput transcriptome analysis performed previously [49]. We studied the levels of expression of 8 transcripts including: the intermediate filaments vimentin (aloalleles a and b) and nestin, the ECM components versican, tenascin-C, fibronectin, collagen type 1 alpha 2, and the enzyme chondroitin 4-sulfotransferase, that is important for the synthesis of CSPGs. Of note, we found that SCI increased significantly the levels of these transcripts at 6 dpt in NR-stage froglets, but not in R-stage animals (Fig. 3W), providing further support to the different glial response in both stages.
In summary, we observed a scar formation in response to injury in NR-stage, which is absent in R-stage. A clear difference in the expression of glial scar makers in response to injury was found in R- and NR-stages. On the one hand, we found a transient increase of vimentin, fibronectin and collagen proteins in R-stages, and no important changes at the RNA levels of glia scar markers. On the contrary, in froglets, a delayed and sustained increase in the protein levels of vimentin, collagen, and CSPG was observed, together with a steady increase of their mRNAs levels.
Characterization of the zGFAP::EGFP transgenic line
For a better understanding of the cellular response triggered by SCI in R- and NR-stages, we decided to prepare a transgenic line that could label most of the cells in the central canal. Although it has been demonstrated that a GFAP gene was lost in Xenopus during anuran evolution [50], we decided to use the zebrafish GFAP (zGFAP) regulatory promoter regions to prepare a transgenic line in Xenopus because of the following reasons: i) based on evolutionary conservation, it is very possible that the main regulators of the gene-regulatory networks operating in glial cells are maintained, because of that we hypothesized that the regulatory promoter regions of the zebrafish GFAP gene could drive expression of a transgene in the same cells in which it is active in zebrafish; ii) a transgenic line using a 11.6 kb region of zGFAP regulatory sequences had been reported and showed proper expression of the transgene in glial and neural progenitor cells in zebrafish [51], and iii) GFAP is usually expressed in many of the cells that are present in the CC including among others radial glial cells, neural stem, neural progenitors, astrocytes and ependymal cells [52].
Before the generation of the transgenic line, and to test our assumption that the zGFAP promoter will drive proper expression in X. laevis spinal cord, we electroporated the spinal cord with a construct in which EGFP expression is driven by the zGFAP regulatory sequences (zGFAP::EGFP), or a control transgene driven the expression of EGFP under a constitutive active promoter (CAG::EGFP). Electroporation of CAG::EGFP revealed an abundant and ubiquitous expression in most of the cells of the spinal cord (see supplementary material, Fig. S3A-C); compared to a more specific and selective expression after electroporation of the zGFAP::EGFP construct, which labeled a specific group of cells in the spinal cord that have a radial glial cell morphology (see supplementary material, Fig. S3D-F). This analysis suggests that the zebrafish regulatory regions drive proper expression in Xenopus. Because of this, we prepared a transgenic line in X. laevis using the same genomic region from the zGFAP described before [51]. The line obtained was named Xla.Tg(Dre.gfap:EGFP)Larra, for short zGFAP::EGFP.
EGFP expression in the transgenic line was detected in the central nervous system (CNS) throughout development (see supplementary material, Fig. S3G-J’). At NF stage 43 and 50 the transgene was expressed in the retina, tectum, cerebellum and spinal cord, but not in the more anterior region of the CNS (Fig. 4A-C). Coronal sections of the cervical, thoracic and lumbar spinal cord showed expression of EGFP in cells, mainly in the dorsal portion of the spinal cord, that have a radial glia morphology, with their apical surface lining the central canal and a long projection to the meningeal layer (Fig. 4D and see supplementary material, Fig. S3K). Most EGFP+ cells also expressed Sox2, a marker of neural stem progenitor cells (NSPC), but not all Sox2+ cells expressed EGFP (Fig. 4E-F’’ and see supplementary material, Fig. S3L-M’’). We noted that the region of the spinal cord with cells expressing EGFP corresponds to the same region containing cells that are actively proliferating in uninjured animals, as demonstrated before by the incorporation of thymidine analogues [47–48].
EGFP+ cells were also found in the spinal cord in NF stage 66 froglets, but have a very different shape and distribution. At this stage only a reduced group of EGFP+ cells are in contact with the CC, mainly on the most dorsal portion (Fig. 4G and I), and extend a dense array of projections towards the meningeal layer (Fig. 4G, I and I’). The most abundant cells expressing EGFP correspond to cells that are not in contact with the CC, but also have cellular projections (Fig. 4G, I, I’’ arrowheads). As shown previously, low levels of Sox2 expression were detected in cells lining the central canal, particularly in the subependymal layer co-localizing with EGFP (Fig. 4H,I’’). Most EGFP+ cells at NF stage 66 co-expressed the Brain Lipid Binding Protein (BLBP) (Fig. 4J-L’), and glutamine synthase (GS) (Fig. 4M-O’), two markers of radial glial cells during early development, and markers of astrocytes at later stages. Base on their morphology, location, and co-expression of other markers we propose that at NF stage 66 most EGFP+ cells at the subependymal layer correspond to astrocytes.
For a more accurate identification and morphological characterization of the cells expressing EGFP in R-stage, we carried out immunogold staining using anti-EGFP antibodies. Gold particles were found in the cytoplasm of cells that are in contact with the central canal (Fig. 4P) and contain intermediate filaments (Fig. 4P’,P’’). Gold particles were also found on cellular projections that were in direct contact with blood vessels on the meningeal side of the spinal cord (Fig. 4Q). Based on their morphology, co-expression of Sox2 and proliferative activity we envision that most EGFP+ cells in the spinal cord of R-stage animals correspond to NSPC with a radial glial morphology. According to our previous characterization of the cells lining the central canal, EGFP+ cells in the R-stage correspond to cells type II and III and is almost not detected in froglets [48].
To unequivocally address the identity of EGFP+ cells in the zGFAP::EGFP transgenic animals at R-stage (NF stage 50) we separated EGFP+ and EGFP− cells using fluorescent activated cell sorter (FACS), and performed RNAseq in each population. Total RNA from EGFP+ and EGFP− cells was sequenced, and the reads mapped. Bioinformatics analysis (see supplementary material Fig. S4A) identified 1,718 transcripts with different levels of expression between EGFP+ and EGFP− cells, 147 of them enriched in EGFP+ cells, including EGFP with the highest fold of change, and 1,571 that showed lower levels of expression in EGFP+ cells (see supplementary material, Fig. S4B). Importantly, gene ontology analysis of the genes enriched in EGFP+ cells showed that the two main categories of biological processes enriched correspond to neural precursor cell identity and stem cell proliferation categories (Fig. 4R). Furthermore, a cluster dendrogram analysis comparing the gene profile of EGFP+ and EGFP− cells with a database of different cells types of the CNS [53] revealed a close correlation between EGFP+ cells and astrocytes, that probably include a radial glial cell signature (Fig. 4S). This molecular analysis confirmed that most of the EGFP+ cells in the zGFAP::EGFP transgenic line correspond to NSPC at NF stage 50, and validated these transgenic animals as a bona fide tool to study the response of NSPC to SCI.
In summary, the use of the zebrafish regulatory regions of GFAP allowed the generation of a X. laevis transgenic line in which EGFP expression recapitulates the expected pattern in the CNS. In R-stages, it is expressed mainly in NSPC with a radial morphology, and later in NR-stages is found primarily in astrocytes, providing a useful and reliable tool to study and characterize the function of these cells in different developmental and regenerative contexts.
Response of neural stem progenitor cells to spinal cord injury
We used the transgenic line described above to study the response, and function of NSPC after SCI in R-stage animals. To evaluate the proliferative response of these cells, transected and sham controls animals were incubated with a pulse of 5-Ethynyl-2´-deoxyuridine (EdU) between 20 and 36 hours after injury (Fig. 5A). Low levels of EdU incorporation in EGFP+ cells were observed in sham operated animals (Fig. 5B,C). Contrary to that, a massive proliferation of EGFP+ cells was observed after SCI (Fig. 5B,C). As a control, EdU+ cells were counted in the intestine, an organ with a high proliferation rate, and no change on the proliferative rate was observed (see supplementary material Fig. S5A-C), indicating that the activation of proliferation raised by SCI was specific for NSPC in the spinal cord. These results are very similar to those reported for the activation of Sox2+ cells in R-stage [47] giving further support to the finding that most EGFP+ cells at this stage co-express Sox2.
To study in detail the response of EGFP+ cells, we fixed animals at different days, and performed immunofluorescence against EGFP in longitudinal sections. To allow a more detailed analysis of the cellular responses in the ablation gap, we performed a resection of the spinal cord that implicates the complete removal of a piece of the spinal cord of approximately 200 µm. In uninjured animals, but now in a longitudinal section, the EGFP+ cells showed their radial morphology and its dorsal and lateral location (Fig. 5D-5D’’). At 2 days post resection (dpr), both ends of the spinal cord were approximately 200 µm apart (Fig. 5E), and many round shaped EGFP+ cells were found at the edges of the rostral and caudal stumps (Fig. 5E’,E’’, arrowheads). Interestingly, at 6 dpr, EGFP+ cellular processes started to extend from the rostral and caudal stumps towards the ablation gap (Fig. 5F-F’’, arrowheads). At 7 dpr some of these processes were even able to cross the complete ablation gap (Fig. 5G,G’, arrowheads). At 10 dpr, EGFP+ cells populated the injured site (Fig. 5H,H’) and some reorganization of the central canal is observed (Fig. 5H’). At 20 dpr, some recovery of the anatomy of the spinal cord was observed (Fig. 1T, and [46–47]). EGFP+ cells were starting to acquire their normal location; however, a radial glial morphology was not observed, and these cells were now present in the ventral level (Fig. 5I-I’’).
Regarding the cellular processes extending into the ablation gap observed at 6–7 dpr we hypothesized two alternatives: i) they could correspond to glial extensions that can provide a substrate for axon growth and pathfinding, something that has been proposed before [42] or ii) in agreement with our previous findings on the role of neurogenesis on spinal cord regeneration [47], these processes could be axons from the new neurons generated from the EGFP+ cells, that because of the half-life of the EGFP protein allowed the study of the cell fate of the EGFP+ cells. To analyze these two possibilities, we performed immunofluorescence against acetylated tubulin, which labels axons and cilia, in the same time points depicted above. As expected, in uninjured animals, acetylated tubulin does not co-localized with EGFP in axons (Fig. 5J-J’’, see arrowheads), but there is co-localization in cells in the central canal probably because acetylated tubulin is present in ciliated cells (Fig. 5J-J’’, see arrows). However, at two days after injury a clear co-localization of acetylated tubulin with EGFP was detected, primarily at the edge of the stumps in structures that are reminiscent of axons and axon growth cones (Fig. 5K-K’’, arrowheads). Importantly, at 6 and 7 dpr most of the EGFP+ cellular processes extending into the ablation gap co-localized with acetylated tubulin (Fig. 5L-L’’,M-M’’). Something similar was observed at 10 dpr (Fig. 5N-N’’). An expression pattern of acetylated tubulin like the uninjured spinal cord is observed at 20 dpr, however, some co-localization of acetylated tubulin and EGFP was still observed in the axonal tracts (Fig. 5O-O’’, see arrowheads).
These results support the hypothesis that the EGFP+ cellular processes crossing the ablation gap correspond to axons because of their morphology, and the co-expression of acetylated tubulin. The fact that they are EGFP+ indicates that most probably they correspond to new neurons formed from the NSPC present in the central canal.
Fate and function of neural stem progenitor cells in spinal cord regeneration
To further evaluate the fate of NSPC, we took advantage of the persistence of EGFP expression. EGFP+ and EGFP− cells were isolated by FACS before and after injury, and the expression levels of the following markers were measured by RT-qPCR (Fig. 6A): i) sox2 and nestin, for NSPC; ii) neurogenin3, achaete-scute homolog 1 (Ascl1), neurogenin2a, doublecortin (Dcx), and neuroD1, for neuronal precursors and neurogenic differentiation; (iii) aldehyde dehydrogenase1l1 (aldh1l1) and vimentin-a, for astrocytes; and the myelin binding protein (Mbp), and Sox10 for oligodendrocytes.
An increase of approximately 450 and 130 times, was observed in the ratio of EGFP levels between EGFP+ and EGFP−cells, at 2 and 6 dpt, respectively. These ratios were many times higher than the ratios detected in uninjured animals (Fig. 6B), probably explained by the increase on the total number of EGFP+/NSPC, because of its massive proliferation induced by SCI (Fig. 5C, and [47]). Supporting the increase in the proliferation of NSPC induced by transection, we observed higher ratios of sox2 and nestin, between EGFP+ and EGFP− cells, at 2 and 6 dpt (Fig. 6C,D). The most probable explanation for this rise is an increase in the number of EGFP+ cells; however, we cannot discard the possibility that the higher ratios were also explained by an increased expression of these genes in each EGFP+ cell.
Interestingly, the early neurogenic markers Ascl1, Neurogenin2a, Neurogenin3, NeuroD1 and Dcx were also increased at 2 dpt, and in some cases also at 6 dpt (Fig. 6E-I). A similar response was observed in astrocytes marker such as vimentin-a and Aldh1l1 (Fig. 6J,K). As an indication that the transgene zGFAP::EGFP is not expressed in oligodendrocytes, lower levels of Sox10 and Mbp were detected in EGFP+ than EGFP− cells, in uninjured animals, and these levels were even smaller at 2 dpt, probably as a consequence of the enrichment in the neuronal, and astrocytic lineage (Fig. 6L,M). In line with the analysis depicted above, these results showed that SCI activates NSPC proliferation, followed by the fate of this neural precursor to the neurogenic and/or astrocytic lineage, but not to oligodendrocytes.
To study the function of NSPC we prepare a transgenic line with the nitroreductase/metronidazol (NTR/MTZ) system in order to specifically ablate these cells [54]. Spinal cord electroporation with a zGFAP::mCherry-NTR construct (see supplementary material, Fig. S6A) followed by incubation with 10 mM MTZ or vehicle (see supplementary material, Fig. S6B) showed a very effective ablation of mCherry+ cells once animals were treated with MTZ compared with vehicle treatment (see supplementary material, Fig.S6C-R). Based on these results we prepared the transgenic line Xla.Tg(Dre.gfap:mCherry-Nitroreductase)Larra (see supplementary material, Fig.S5S-U), for short zGFAP::mCherry-NTR.
We use this line to evaluate the effects of NSPC ablation in the ability of R-stage animals to regenerate the spinal cord and recover their swimming ability. For this, four groups of animals were considered: i) sham operated animals treated with vehicle or MTZ, and ii) resected animals treated with vehicle or MTZ. Animals were incubated with vehicle or MTZ for 1 week before sham operation or spinal cord transection (Fig. 7A). Efficient ablation of mCherry+ cells was attained at 7 days after incubation with MTZ (Fig. 7B-G). After transection, we measured the swimming ability of each group. Treatment with MTZ has no effect on the ability of sham-operated animals to maintain their swimming ability compared to vehicle treated animals (Fig. 7H, and data not shown). Importantly, at 15 and 25 dpr animals incubated with MTZ showed a diminished swimming ability compared to vehicle treated siblings (Fig. 7H, compare red and green boxes). In addition, we performed Sox2 immunostaining in sections of the spinal cord at 30 dpt, and found that treatment with MTZ, as expected, resulted in a strong reduction of Sox2+ cells precluding the proper regeneration of the spinal (Fig. 7I-L). These results indicate that NSPC cells are necessary for regeneration of the spinal cord of NF stage 50.