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Jul 24, 2012 - system during salamander tail regeneration. Levan Mchedlishvilia,b,1,2, Vladimir Mazurova,1, Kathrin S. Grassmea,3, Kerstin Goehlera,b, ...
Reconstitution of the central and peripheral nervous system during salamander tail regeneration Levan Mchedlishvilia,b,1,2, Vladimir Mazurova,1, Kathrin S. Grassmea,3, Kerstin Goehlera,b, Bernhard Robla,b, Akira Tazakia,b, Kathleen Roenscha,b, Annett Duemmlera,b, and Elly M. Tanakaa,b,4 a Max Planck Institute of Molecular Cell Biology and Genetics, 01307 Dresden, Germany; and bDeutsche Forschungsgemeinschaft Center for Regenerative Therapies Dresden, Cluster of Excellence, University of Technology Dresden, 01307 Dresden, Germany

Edited by Anders Bjorklund, Lund University, Lund, Sweden, and approved July 6, 2012 (received for review October 12, 2011)

We show that after tail amputation in Ambystoma mexicanum (Axolotl) the correct number and spacing of dorsal root ganglia are regenerated. By transplantation of spinal cord tissue and nonclonal neurospheres, we show that the central spinal cord represents a source of peripheral nervous system cells. Interestingly, melanophores migrate from preexisting precursors in the skin. Finally, we demonstrate that implantation of a clonally derived spinal cord neurosphere can result in reconstitution of all examined cell types in the regenerating central spinal cord, suggesting derivation of a cell with spinal cord stem cell properties. neural stem cell

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egeneration of the central body axis occurs after tail amputation in salamander amphibians. During this process the spinal cord regrows, and the correct number of segmented vertebrae and myotomes are formed (1). Additionally, neural crest derivatives, such as melanophores, and the peripheral nervous system (PNS), including dorsal root ganglia (DRG) and Schwann cells, are regenerated (2–4). An important challenge is to define and study the stem cells that are responsible for regenerating the CNS and PNS. Previously, we used electroporation of GFP expression plasmids into the spinal cord to identify and track the radial glial cells that contribute to regenerating the spinal cord in the salamander Ambystoma mexicanum (axolotl) (5, 6). Live cell tracking showed that single cells could give rise to clones populating multiple molecular domains of the regenerating spinal cord (6). These results indicated that multipotent progenitors exist during spinal cord regeneration. Because of the transient expression of the plasmids, the long-term fate of the stem cells could not be tracked, so the origin of the PNS was not addressed. The source of neural crest derivatives in the regenerated tail has been an open question since 1885 (7). During early development the neural crest arises in the dorsal region of the neural tube, from which cells emigrate to the surroundings to form neurons and glia of PNS, smooth muscles, head skeletal elements, enteric neurons, and melanophores (8). It is not yet known conclusively, however, whether during regeneration newly regenerated neural crest structures derive from a population of neural crest-like cells that migrate out of the regenerating spinal cord or arise directly from cells in the periphery. Immunohistochemical studies using markers such as HNK1 suggested that there may be a population of cells in the lateral walls of the spinal cord with neural crest properties (4). Furthermore, morphological studies suggested that cells may migrate via the forming ventral roots to populate the spinal ganglia outside the spinal cord. Such findings could be consistent with recent findings in mouse that boundary cap cells can act as a neural crest source (9). To track the origin of neural crest structures during newt tail regeneration, Benraiss et al. (3) attempted to label spinal cord cells via biolistic transfection of a human alkaline phosphatase (AP) expression vector. In such studies, they later observed AP expression in melanophores and Schwann cells of the periphery but not in DRG. However, because the AP gene

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was driven by the ubiquitous SV40 promoter, it was impossible to exclude the possibility that transfection of the cells outside the spinal cord had occurred also. Here we examine the regeneration of CNS and PNS during axolotl tail regeneration. By transgenic labeling of tissues we show that DRG and Schwann cells derive from cell pools associated with the central regenerating spinal cord. In support of a central source of PNS regeneration, we describe generation of neurosphere cultures from the axolotl spinal cord that, when engrafted back into the animal, integrate and undergo extensive self-renewal before forming significant portions of the regenerated spinal cord and PNS. Clonal GFP+ neurospheres engrafted and contributed to all regions of the central spinal cord, indicating the ability to derive a spinal cord neural stem cell in clonal culture. Results Normal Number and Spacing of DRG Are Reconstituted During Axolotl Tail Regeneration. In axolotls, the regenerating tail ultimately

achieves overall dimensions comparable to control, unamputated tails. Previous literature established that DRG were regenerated, but there was no demonstration that the correct number and spacing of DRG were reestablished after tail amputation. To assess whether the correct complement of DRG was regenerated, we used whole-mount staining with NeuN and βIII-tubulin immunofluorescence to visualize the tail DRG. DRG were identifiable as bright clusters next to the spinal cord (Fig. 1, arrowheads), as confirmed by cross-sectioning the whole-mount samples. Tails were amputated at the 15th myotome, and the number and spacing of regenerated DRG were determined at 35 d postamputation (dpa), when the regenerated tails first achieved an overall length comparable to uncut controls, and at 49 dpa (Fig. S1 and Table S1). At 35 dpa, we observed that the number of regenerated DRG was comparable to control samples (Fig. 1E). However, the spacing between the DRG at myotome level 22–28 was significantly smaller in the regenerating samples than in controls (P < 0.0001). By 49 dpa the inter-DRG spacing in the regenerated samples was similar to that in controls (Fig.

Author contributions: L.M., V.M., and E.M.T. designed research; L.M., V.M., K.S.G., K.G., B.R., and K.R. performed research; A.T., K.R., and A.D. contributed new reagents/analytic tools; L.M., V.M., K.S.G., and E.M.T. analyzed data; and L.M. and E.M.T. wrote the paper. The authors declare no conflict of interest. This article is a PNAS Direct Submission. Freely available online through the PNAS open access option. 1

L.M. and V.M. contributed equally to this work.

2

Present address: Department of Physiological Genomics, Institute of Physiology, Ludwig Maximilian University of Munich, 80336 Munich, Germany.

3

Present address: University of Muenster, Angiogenesis Laboratory, Röntgenstr. 20, 48149 Muenster, Germany.

4

To whom correspondence should be addressed: E-mail: [email protected].

See Author Summary on page 13477 (volume 109, number 34). This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10. 1073/pnas.1116738109/-/DCSupplemental.

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Fig. 1. DRG number is restored by 35 d, but the correct inter-DRG distance is restored by 49 d of regeneration. Axolotl tail tissue was immunostained simultaneously using mouse anti–βIII-tubulin and mouse anti-NeuN antibody and then was detected in a single channel using a Cy5-coupled anti-mouse antibody. (A) Merged DIC/fluorescence image The arrow shows the amputation plane. (Scale bar: 1 mm.) (B–D) Bright cell clusters adjacent to the spinal cord can be observed at DRG 11–13 (B, B′, arrowheads) and 17–19 (C, C′, arrowheads). Postcloaca DRG 24–26 in the posterior tail region can identified by the axonal route emerging from the spinal cord region (D, D′, arrowheads). (Scale bars: B′–D′ and B′′–D′′, 0.7 mm.) (E) Inter-DRG distance was measured in all DRG posterior to the cut plane in amputated animals and compared with comparable DRG in control animals at 35 dpa. The inter-DRG distance is smaller in regenerated tails than in controls when assessed by the Student’s t test (P < 0.0001). (F) At 49 dpa the inter-DRG distances are comparable in regenerated tails and control animals (P = 0.39 for difference, Student’s t test).

1F) (P = 0.39 for difference, Student’s t test). At 35 dpa a DRGfree zone still remains in the posterior tip of the tail. Between 35 and 49 dpa, the myotome size in the posterior half of the tail increases, and the DRG-free regions shrink, resulting in comparable inter-DRG spacing in regenerated and control tails. These data show that during axolotl tail regeneration, a normal complement and spacing of DRG are reconstituted but that the number of regenerated DRG is restored before the final spacing is achieved. Spinal Cord Transplantations Suggest That Newly Formed DRG Can Derive from the Spinal Cord Region. The newly formed DRG could

derive from several likely sources. First, it is possible that the regenerating spinal cord harbors a neural crest-like population or boundary cap cells that, as in embryonic development, emigrate from the tube to generate neural crest structures (8–10). The expression of neural crest-associated transcription factors such as MSX1 and PAX7 in the dorsal region of the regenerating spinal cord suggests that neural crest may be reformed during Mchedlishvili et al.

spinal cord regeneration (6, 11). Second, it is possible that neural crest precursors residing outside the spinal cord migrate posteriorly to generate the new DRG. To determine whether the regenerating spinal cord represents a potential source of cells for the new neural crest-derived structures, we labeled the spinal cord by transplanting a 2- to 4-mm segment of spinal cord encompassing the 16th myotome level from GFP-expressing transgenic animals (6, 12) into white host animals (n = 8) and 10 d after healing amputated the tail at myotome 15, leaving 2 mm of GFP+ spinal cord (Fig. 2). The tails were allowed to regenerate for 15 wk before analysis (Fig. 2 A–C). Three regenerated tails were cryosectioned and stained for the marker βIIItubulin, which highlights the DRG. The proportion of GFP+ DRG cells was analyzed starting from myotome 22 to ensure that the DRG being analyzed represent regenerated DRG (Fig. 2 D– F, arrows). In the first tail, 21 DRG (11 left, 10 right) were found, of which 14 were 100% GFP+. In the rostral portion three DRG were completely GFP−, and four were mosaically GFP+/−. In tail two, 20 DRG (11 left, 9 right) were regenerated, of which 16 PNAS | Published online July 24, 2012 | E2259

Fig. 2. Transplanted GFP+ spinal cord participated in regeneration of spinal cord and DRG. A 2- to 4-mm gap in the spinal cord was made in the axolotl tail and was replaced with a comparable length of spinal cord from the germline GFP+ transgenic animal. (A) Image immediately after the transplanted tail was amputated. (B) At 29 dpa the tail has regenerated much of its length, and the regenerated spinal cord is largely GFP+. (C) GFP+ spinal cord regenerates 105 dpa. Cross-sections from spinal cord-transplanted animal immunostained for βIII-tubulin. (D–F) Three examples of DRG adjacent to transplanted, regenerated GFP+ spinal cord. (D) An example in which the whole DRG was GFP−. (E) An example of a DRG with a mixture of GFP+ and GFP− cells. (F) An example in which the whole DRG was GFP+. In D–F, GFP is shown as green, βIII-tubulin as red, and Hoechst staining as blue. (Scale bars: A–C, 1 mm; D, F, 100 μm; E 50 μm.)

were completely GFP+, two were GFP−, and two were GFP+/− at myotomes 22–24. In the third tail, 17 regenerating DRG were found (5 right and 12 left), of which 16 were GFP+ and one was GFP−. In previous work examining the origin of the central spinal cord, spinal cord transplantation indicated that a 500-μm zone represented the source zone for spinal cord regeneration (6). We also attempted to analyze spinal cord transplantations in which a 500-μm GFP+ zone remained after tail amputation for DRG. Such transplantations result in regenerates in which, although there are GFP+ cells all along the regenerating spinal cord, the spinal cord in some cases is a mosaic of GFP+ and GFP− cells because the junction of cell healing between host and donor is irregular. Therefore, the percentage of GFP+ DRG was highly variable. In tail one, 13 DRG were found next to the GFP+ spinal cord (six left and seven right), four of which were mosaic GFP+/−, and nine of which were GFP−. In tail two, 18 DRG were found (11 left and 7 right), of which the two most rostral DRG were GFP−, the six following caudal DRG were GFP+/−, and the 10 most caudal DRG were GFP+. In the third tail, 15 DRG were found (eight left and seven right), all of which were GFP−. In this tail the spinal cord was constructed mosaically (GFP+/−); approximately one-fourth of the spinal cord cells in the cross-section were GFP−. Taken together, however, these results indicate that a population of cells associated with the central spinal cord can be a significant source of cells for the DRG and PNS. Melanophores Can Derive from Peripheral Structures. It was shown recently that, during development, melanocytes, another neural crest derivative, arise from precursors residing in peripheral nerve E2260 | www.pnas.org/cgi/doi/10.1073/pnas.1116738109

fibers (13). Furthermore, in Xenopus, studies of neural crest-ablated tadpoles showed that the melanophores regenerate from preexisting unpigmented melanophore precursors in the skin adjacent to the amputation site (14). This mechanism is similar to that described for the zebrafish fin regeneration, in which melanoblasts in the skin generate the new melanophores in the regenerated fin (15). To test whether precursors in axolotl skin contribute pigmented cells to axolotl tail regeneration, we transplanted skin from wild-type animals into the white mutant hosts that have no mature melanophores in the skin because the melanophore precursors do not migrate from the neural crest region during development (16). After healing for 4 d, the tails were amputated through the graft. After 26 d of regeneration, cells from the wild-type skin patch had clearly formed melanophores that populated the regenerated tail. (Fig. S2). This result indicates that a significant portion of the melanophores in the regenerated axolotl tail probably arise from peripherally located precursor cells. Because of the technical difficulty of detecting GFP in pigmented melanophores, we could not address whether the spinal cord also is a source of melanophores in regenerated tail. Generation and Characterization of Axolotl Neurospheres as a Transplantable Cell Source for CNS and PNS Regeneration. To refine our

analysis further, we endeavored to isolate and implant GFP+ spinal cord stem cells into the spinal cord to track the source of PNS. We used culture conditions previously described for mammalian neurospheres to derive cultures from the GFP+ axolotl spinal cords (17). Within 2 wk, cell clusters resembling mammalian neurospheres emerged in culture (Fig. 3 A–C). To promote the formation of primary neurospheres, we tested dif-

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ferent peptide growth factors reported to support mouse primitive, embryonic, or adult neural stem cells and found that the addition of 20 ng/mL FGF produced the highest yield of neurospheres, with an average of 215 neurospheres per 50,000 seeded cells (Fig. 3D) (17–19). We characterized the cultured neurospheres for marker expression, their ability to form differentiated neural cells in culture, and their ability to reconstitute neural cell types upon implantation into the spinal cord. We first determined whether the axolotl neurospheres contained cells expressing progenitor cell markers and/or differentiated cell markers. We found that the majority of cells in the neurosphere were positive for Sox2, which marks the neural progenitor cell pools in the vertebrate CNS (Fig. 3 E and I). Consistent with a stem/progenitor cell identity of neurosphere cells, Musashi-1 (20), which stains the ependymoglial cells of the axolotl spinal cord, was found in the majority of neurosphere cells (Fig. 3 F and J). The neurospheres also stained strongly for GFAP, a marker for radial glial cells and astrocytes (Fig. 3 G and K). Because the mature and regenerating axolotl spinal cord consists largely of radial glial-like cells, we interpret the GFAP+ cells in the neurospheres to be radial glia with neurogenic capacity. Finally, staining with the neuronal Mchedlishvili et al.

marker βIII-tubulin showed the presence of differentiated neurons in the periphery of the neurospheres (Fig. 3 H and L). To test the in vitro differentiation potential, the nonclonal neurospheres were plated onto poly-L-lysine/laminin–coated surfaces under growth factor-poor conditions, where they displayed robust outgrowth of βIII-tubulin+ axons but no myelin basic protein-positive (MBP+) oligodendrocytes (Fig. 3 M and O, Inset). Under these conditions GFAP staining was visible but attenuated compared with floating spheres (Fig. 3N). In contrast, when the cells were exposed to 50 ng/mL Sonic hedgehog (Shh) agonist and 25 ng/mL PDGF, we observed strong staining for the glial marker MBP (Fig. 3O). Implanted GFP+ Neurospheres Integrate into the Spinal Cord and Regenerate Substantial Regions of the Spinal Cord and DRG in Vivo.

We wanted to test the competence of the primary, nonclonal neurospheres to integrate and regenerate the spinal cord and PNS. To achieve integration of the cultured neurospheres into spinal cord tissue, we removed a 1- to 3-mm section of the spinal cord in the tail and placed one or several GFP neurospheres into the gap (Fig. 4A). The animals were allowed to heal (Fig. 4B). To examine if the implanted neurosphere cells had integrated into the injured spinal cord, we took cross-sections of the tail through the engrafted PNAS | Published online July 24, 2012 | E2261

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Fig. 3. Spinal cord neurospheres can be derived in FGF-2 and express neural stem cell markers. (A) Overview of procedure. Spinal cord tissue was isolated from GFP+ axolotl tail tissue, dissociated in papain, and expanded in serum-free medium containing growth factors and supplements (B27). (B) Low-magnification view of neurosphere culture after 2 wk. (Scale bar: 200 μm.) (C) Higher-magnification view of neurosphere. (Scale bar: 100 μm.). (D) Growth-factor dependence of neurosphere derivation. After dissociation, axolotl spinal cord cells were counted and expanded in serum-free medium in the presence of various growth factors; 50,000 cells were plated for each condition. Among the different factors and combinations of factors [e.g., FGF-2, EGF, FGF-2/EGF, PDGF, and leukemia-inhibitory factor (LIF)], FGF-2 yielded the highest number of neurospheres. The y-axis represents the yield of neurospheres per 50,000 plated cells. n = 3. (E) Confocal imaging of anti-Sox2 of a cross-section of an immunostained axolotl spinal cord shows neural progenitor cells. (F) Anti– Musashi-1 immunostaining of a spinal cord cross-section corroborates luminal location of neural progenitor cells. (G) Anti-GFAP immunostaining of a spinal cord cross-section reveals the presence of radial glia in the axolotl spinal cord. (H) Anti–βIII-tubulin immunostaining of a spinal cord cross-section shows neuronal cells in the axolotl spinal cord. (I) Confocal imaging of an anti-Sox2–immunostained neurosphere cross-section shows that the majority of cells are neural progenitors. (J) Anti–Musashi-1 immunostaining of a neurosphere cross-section shows that many cells are neural progenitors. (K) Anti-GFAP immunostaining of a neurosphere cross-section indicates that the majority of cells are apparently GFAP+. (L) Anti–βIII-tubulin immunostaining of a neurosphere indicates that some cells are or have differentiated into neuronal cells. Red indicates the antigen-specific immunostaining signal; blue indicates Hoechst staining. (Scale bars: E–L, 20 μm.) (M–O) To induce neuron outgrowth, neurospheres were plated onto poly-L-lysine/laminin–coated Petri dishes. After 7 d in culture, neurosphere-derived cells were positive for βIII-tubulin immunostaining (M) and were faintly positive for GFAP (N) but were negative for MBP immunostaining (O, Inset). Data shown in M, N, and O Inset are representative of three independent experiments. (O) In the presence of Shh agonist (50 ng/ mL) and PDGF (25 ng/mL), neurosphere cells differentiated into MBP+ glia. Data shown are the results of two independent experiments. Inset shows MBP staining of a neurosphere grown in the absence of Shh agonist and PDGF. (Scale bars: M, 50 μm; N and O, 100 μm; O Inset, 50 μm.)

Fig. 4. GFP+ cells derived from a passaged neurosphere implant contribute to peripheral nerves and DRG. (A) After the removal of an ∼1-mm section of spinal cord, the neurospheres were implanted in the lesion. (B) After 1–2 wk of wound healing, the implanted neurospheres were integrated in the spinal cord tissue. The tail was amputated close to the integrated GFP+ cells, maintaining them within the 500-μm zone. (C) The integrated GFP+ cells contribute to spinal cord regeneration upon amputation. (D) MBP immunostaining of a tail cross-section containing regenerated EGFP+ spinal cord derived from the passaged neurosphere implant. An MBP+/EGFP+ nerve is shown in the box. (E) Confocal image of peripheral nerve showing GFP+ nuclei and MBP+/GFP+ myelin. (F) An anti-MBP–stained nerve myelin sheet. (G) GFP channel view of the nerve indicates EGFP+ cell nuclei and myelin cytoplasm. Green indicates EGFP+ cells, red indicates MBP immunostaining, and blue indicates Hoechst staining. (H) βIII-Tubulin immunostaining of the same cross-section. A DRG containing βIIItubulin+/GFP+ cells is shown in the box. (I–K) Confocal image of DRG showing βIII-tubulin+/GFP+ cell and nerves. Green indicates EGFP+ cell bodies and nerves; red indicates βIII-tubulin immunostaining; blue indicates Hoechst staining. (Scale bars: D and H, 50 μm; E, F, and I–K, 10 μm.)

neurospheres at different time points (4, 7, 14, and 21d after implantation) and immunostained the sections for GFAP (Fig. S3). At day 4 the engrafted neurosphere cells made a continuous mass with the endogenous radial glial cell layer but formed an ectopic protrusion from the spinal cord tube (Fig. S3A). At day 7 the epithelial cell contact between the grafted neurosphere and the spinal cord was more pronounced (Fig. S3B). Again, GFAP immunostaining seemed reduced in most of the GFP+ cells. By 14 d postimplantation, the cells had integrated into the structure of the host spinal cord (Fig. S3C). By day 21, and later, cells could be seen clearly in the neuronal layers, and GFP axonal projections also could be seen (Fig. S3D). To examine whether neurosphere cells participate in regeneration, the tail was cut adjacent to the implanted cells, and the distribution of the GFP+ cells was followed in vivo (Fig. S4). In 7 of 12 implantations GFP cells were distributed along the whole spinal cord. The example in Fig. S4 was reamputated three times in succession. Each time the GFP cells were found all along the length of the spinal cord, demonstrating the ability of the implanted cells to expand extensively. Cross-sections of the regenerating spinal cord showed that the GFP cells had solidly populated large regions of the spinal cord, as in the half of the spinal cord shown in Fig. S4 G–I. Colocalization of GFP with cell type-specific markers indicated that the integrated cells formed GFAP+ radial glia (Fig. S4G), MAP2+ neurites (Fig. S4J), and NeuN+ neurons (Fig. S4M). To examine if the GFP+ neuronal cells form motor neurons with connections to muscle, we retrogradely labeled motor neurons by injecting tail muscles with rhodamine dextran and fixed the tails 24 h later. Cryosectioned samples show that a cell body in the neural layer of the ventral spinal cord was labeled consistent with a motor neuron identity (Fig. S5 A–D, arrows). The labeled cells expressed the neural-specific marker NeuN E2262 | www.pnas.org/cgi/doi/10.1073/pnas.1116738109

(Fig. S5 A–D, arrows), indicating that primary neurospheres elaborate neurons that make connections with the periphery. Finally we examined whether the implanted neurospheres acquired markers of positional identity upon implantation. Because tail cutting requires cells to form more posterior structures, we examined the expression status of posterior Hox proteins A9 and A13 during spinal cord regeneration using polyclonal antibodies (Materials and Methods). In mature portions of the spinal cord we observed HoxA9 expression in a subset of cells in the outer, neuron-containing layer of the spinal cord, but cells close to the lumen were negative (Fig. S6 G and H). We observed no HoxA13 expression in the mature part of the spinal cord (Fig. S6 Q and R). Both HoxA9 and HoxA13 expression was observed in the 6-d regenerating spinal cord, with uniform, nuclear expression found toward the caudal regions of the regenerate (Fig. S6 A, B, D, E, K, L, N, and O). Before implantation, the axolotl neurospheres expressed nuclear HoxA9 protein in some peripheral cells of the neurosphere. In contrast, HoxA13 showed diffuse cytoplasmic expression. After implantation of GFP+ neurospheres, examination of 6-d regenerates by confocal microscopy showed that the implanted GFP+ neurosphere cells expressed HoxA9 and HoxA13 in a manner similar to their endogenous neighbors, at different levels along the rostral to caudal axis (Fig. S6 A–F and K–P). These results suggest that the implanted neurosphere cells acquire appropriate rostral/caudal identity profiles in the regenerate. DRG and Schwann Cells Are Formed from Implanted Neurosphere Cells. We next asked whether the implanted cells populate neural

crest lineages. Following the procedure described above, we implanted primary and passaged (P2) GFP+ neurospheres into spinal cord lesions made in the middle of the tail (Fig. 4A). After 7 d of healing, we amputated the tail close to the GFP+ cells and allowed regeneration for 6 wk (Fig. 4 B and C). When we examined Mchedlishvili et al.

Evidence That Clonally Derived Neurospheres Regenerate All Spinal Cord Cell Types. Because neurospheres are widely considered

a means to identify and isolate neural stem cells, we wanted to know whether any cells in our neurospheres had the property of a true multipotent neural stem cell, i.e., the ability to self-renew and to generate all neural cell types of the spinal cord. To address this issue, we developed a clonal neurosphere formation assay for axolotl cells followed after implantation into host tissue (Fig. 5). Because salamander cells do not grow well in isolation, GFP+ cells were cultured together with nontransgenic carrier cells at a ratio of approximately five carrier cells to one GFP+ cell in AggreWell 400 plates (Fig. 5A). The microwells were imaged to record the position of the microwells harboring single GFP+ cells. About 10,000 single GFP+ cells were imaged in different sets of experiments. The vast majority of the cells either died or did not divide during the culturing process. Sixty-seven cells gave rise to neurospheres harboring significant numbers of GFP+ cells. Interestingly, the frequency of successful neurosphere formation corresponds with the frequency of neurosphere formation found under nonclonal conditions (Fig. 3). Neurospheres that were predominantly of GFP+ origin and >60 μm in size (n = 28) were selected for implantation into white hosts with one neurosphere per animal. After 1 wk of healing the tails were amputated close to the GFP+ implant. Three of the 28 clonal neurosphere implants contributed efficiently to the spinal cord regeneration process (Fig. 5 A–F and Figs. S7 and S8). To determine whether GFP+ cells had formed all spinal cord cell types, the caudal part of the regenerate harboring GFP+ spinal cord was amputated and fixed for histological analysis. Crosssections of the tail showed that the entire lateral half of the spinal cord regenerate was repopulated by GFP+ cells (Fig. 5 G–J). Colocalization of GFP with cell type-specific markers indicated that the integrated cells efficiently formed GFAP+ radial glia (Fig. 5G), βIII-tubulin+ neurons (Fig. 5 H and O–R), and MBP+ cells (Fig. 5 O–R, Inset in O and arrowheads). The integrated GFP+ cells also faithfully expressed region-specific transcription factors that specify progenitor cell subtypes such as PAX6 and PAX7 (Fig. 5 I and J). Because in some locations along the anterior/posterior axis, all cells in one lateral half of the spinal cord were GFP+ (Fig. 5G), we determined whether the GFP+ cells deriving from the clone were positive for the ventral-most and dorsal-most cell types. This information indicated that the clonal GFP+ neurosphere cells had contributed daughters to all possible spinal cord cell types. Shh+ and Msx1+ GFP+ floor and roof plate cells, respectively, were found in daughter cells of the implanted GFP+ clonal neurosphere (Fig. 5 K–N, arrows). These results suggested that all spinal cord cell types were produced by the implanted GFP+ clonal neurosphere cells. The tail of the second animal with the integrated, clonally derived neurosphere (Fig. S7) was fixed and analyzed in more depth for the formed neurons and radial glia. As in example one, in some cross-sections the majority of the lateral half of spinal cord was reconstituted from GFP+ cells (Fig. S7G). Cell-specific staining revealed formation of GFP+/NeuN+ neurons (Fig. S7 H and I) and GFP+/GFAP+ radial glia (Fig. S7J). In the third animal (Fig. S8), the implanted neurosphere contributed to Shh+ floor plate cells (Fig. S8 D and G), Pax6+ progenitors (Fig. S8 E and H), and NeuN+ neurons (Fig. S8 F and I) in the spinal cord. In the clonal implantation experiments we did not observe labeled DRG. In sum, our clonal analysis suggests that the diMchedlishvili et al.

Discussion Here we have shown that during axolotl tail regeneration, a faithful complement and periodicity of PNS structures is regenerated. The completeness of PNS regeneration seems to be a unique feature of the axolotl system. Similar tail amputation experiments carried out in Xenopus tadpoles revealed the lack of defined DRG regeneration (14), perhaps because of deficient expression of various embryonic signaling factors in the frog compared with the axolotl (21). Furthermore, through implantation of spinal cord tissue and cultured neurospheres, we show that a substantial proportion of the DRG and the Schwann cells can arise from cells associated with the regenerating spinal cord. Previous immunocytochemical and electron microscopy analysis pointed to lateral exit routes for migratory cells during spinal cord regeneration (2, 4). This feature resembles the late phase of neural crest migration that has been described in the chick (22). In the future, more refined cell tracing of spinal cord subpopulations using genetically marked cells via the cre/lox system in axolotls will allow us to determine the molecular identity, timing, and source of neural crest derivatives in the spinal cord. In contrast, our skin transplantation experiments suggest that preexisting precursors in the periphery contribute significantly to the melanophores in the regenerate. This observation would be consistent with recent lineage-tracing experiments in bird and mouse suggesting that melanocytes derive from precursors in peripheral nerve (13). These observations also would unify the melanophore regeneration mechanisms with zebrafish and Xenopus (14, 23). Because we could not track the spinal cord GFP+ cells into melanophores for technical reasons, we cannot exclude the possibility that spinal cord-derived melanophores might contribute as well. Indeed, the original biolistic transfection-labeling experiments in Pleurodeles waltl suggested that such events may take place, although it was difficult to exclude the possibility that peripheral melanocyte precursors lying close to the spinal cord had been labeled in those experiments (3). As part of our analysis, we have derived a neurosphere-like culture from the axolotl spinal cord that is capable of regenerating substantial parts of the spinal cord and the PNS including the DRG and Schwann cells. The neurospheres are grown under culture conditions similar to those used for mammalian counterparts and differentiate into neuronal and glial cell types under similar induction conditions, supporting the notion that these neuronal and glial cells represent a cognate cell type. We observed generation of motor neurons and sensory neurons from the implanted cells. Furthermore, repeated amputation of the nonclonal neurosphere-implanted tails resulted in at least 106fold expansion of the implanted cells, proving their extensive expansion capacity in vivo. Our clonal neurosphere experiments also suggest the existence in our neurosphere cultures of a cell with the properties of a spinal cord neural stem cell, because we observed engraftment of clonally derived GFP+ cells in all dorsal/ventral cell domains, as analyzed with molecular markers. The term “neural stem cell” often is used in mammalian neurosphere experiments to indicate the ability to give rise to neurons, astrocytes, and oligodendrocytes. Our work demonstrates that clonally derived cells can reconstitute extensively all spinal cord cell types that we could examine in vivo. Although we observed robust contribution to the DRG from nonclonal neurospheres in vivo, we did not observe contribution of the clonally derived neurospheres to DRG in this study. It presently is unresolved whether a neural stem cell that can form both the CNS and PNS exists and simply was not found in the PNAS | Published online July 24, 2012 | E2263

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versity of the central spinal cord cells can be reconstituted from a single cell during regeneration, suggesting multipotency of the cultured axolotl neural stem cells.

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sections containing peripheral nerve, we observed cells with clear nuclear GFP in MBP+ peripheral nerve (Fig. 4 D–G), indicating that Schwann cells can derive from the implanted neurospheres. We often observed DRG that were a mixture of GFP+ and GFP− cells (Fig. 4 H–K), presumably because the GFP+ cells constitute part of the regenerated spinal cord so that both unlabeled and labeled spinal cord cells contribute to the DRG.

Fig. 5. Clonally derived neurosphere efficiently contributes to spinal cord regeneration and reconstitutes the whole complement of spinal cord cell types. (A) GFP+ cells cultured in microwells of an AggreWell 400 plate at clonal density with GFP− white feeder cells. Arrow indicates a single EGFP+ cell in the microwell, imaged on the first day of the culturing procedure. (B) Clonal neurosphere that arose from the single EGFP+ cell indicated by the arrow in A after 34 d of culturing. (C) A clonal GFP+ neurosphere was engrafted into the injured spinal cord of the nontransgenic host. After 1 wk of healing the tail was amputated close to the implant, maintaining GFP+ cells within the 500-μm zone behind the amputation plane. (D–F) Outgrowth of the GFP+ spinal cord from the implanted clonal neurosphere monitored at 7, 24, and 54 dpa. (G–J) Cross-sections of the regenerated spinal cord from F at a caudal level. In this portion of the regenerate the full lateral half of the spinal cord was reconstituted from GFP+ cells. Cross-sections were analyzed by immunofluorescence for expression of cell type-specific markers. (G) Anti-GFAP staining shows GFP+ cells have formed GFAP+ radial glia. (H) Anti–βIII-tubulin staining shows GFP+/βIII-tubulin+ neuronal cells. (I) Cross-section immunostained for Pax6 shows that implanted GFP+ cells express Pax6 in the normal, lateral spinal cord domain. (J) Cross-section immunostained for Pax7 shows that implanted GFP+ cells in the dorsal domain express Pax7. (K and L) Shh in situ hybridization shows clonally derived Shh+/GFP+ floor plate cells (arrows). (M and N) Msx1 in situ hybridization shows Msx1+/GFP+ roof plate cell indicated by arrow. (O) βIII-Tubulin/MBP double-immunostaining of the cross-section. βIII-Tubulin was labeled with Cy5 secondary antibody and is shown in yellow. (P–R) Enlarged views of boxed area in O, which shows the axonal layer of the spinal cord containing MBP+/GFP+ myelin (arrowheads). In G–J, green indicates GFP+ cells; red indicates cell type-specific immunostaining; blue indicates Hoechst staining. In K–M, green indicates GFP, and blue indicates Hoechst staining. In O–R, yellow indicates βIII-tubulin; red indicates MBP; green indicates GFP; and blue indicates Hoechst staining. (Scale bars: A and B, 50 μm; C–F, 1 mm; G–J, 25 μm; K–N, 50 μm; O, 20 μm; P–R, 5 μm.)

clonal studies because of technical limitations or whether a separate pool of stem cells associated with the spinal cord region exists for the CNS and PNS. In the nonclonal neurosphere implants, the number of labeled, implanted cells was much greater E2264 | www.pnas.org/cgi/doi/10.1073/pnas.1116738109

than in the clonal implants, leading to a greater chance that (i) a rare CNS/PNS stem cell was implanted in those studies, and (ii) the implanted CNS/PNS stem cell descendents survived and populated the neural crest-generating zone of the regenerating Mchedlishvili et al.

Materials and Methods Measurement of DRG Regeneration. Animal work was approved by the Saxony State Commission on Animal Welfare. To examine the regenerative potential of DRG during tail regeneration, axolotl larvae of 37.2 ± 0.7-mm length were used. Ten animals were anesthetized in a solution with 0.01% (wt/wt) ethyl-p-aminobenzoate (E-1501; Sigma), and the tails were amputated after the 15th myotome using a sterile scalpel. We measured tail length in the animals at different time points and compared them with an unamputated control cohort (n = 10 for each). The regenerated tails reached a normal tail length, comparable to unamputated controls, by 35 dpa (Table S1). To determine whether the normal number of DRG was regenerated, the number and spacing of DRG were compared in regenerating animals that had been amputated at the 16th myotome posterior to the cloaca (n = 10) and in uninjured control animals (n = 10). We analyzed the number of DRG at two time points after amputation, 35 dpa (n = 5), and 49 dpa (n = 5), to determine whether any further changes in DRG could be observed after the tail had grown to normal length. Tails were fixed with freshly prepared 4% (wt/wt) paraformaldehyde (PFA) before whole-mount immunostaining. All samples were fixed for 12 h at 4 °C. After fixation samples were washed three times for 20 min with PBS. Immunolabeling of Whole-Mount Tails. To analyze the distances between DRG in the regenerated tails and control tails, whole-mount tails were immunostained with NeuN and βIII-tubulin antibodies. Before antibody staining the skin was peeled away from the tails in a Petri dish filled with PBS using scalpel and forceps under a stereo microscope (SZX12-RFA; Olympus). The antibody reaction was done overnight at 4 °C. A Cy5-labeled Fab-fragment goat anti-mouse secondary antibody (Dianova) was used. After staining, the tails were taped onto a coverslip and imaged with in epifluorescence and differential interference contrast (DIC) using a BX61 microscope (Olympus). All images were processed and combined for figures using MetaView, Axiovision 4.6, and Photoshop CS (Adobe). Combined images were used for measuring inter-DRG distances (Photoshop CS; Adobe). Isolation and Culture of Neurospheres. The spinal cords of EGFP transgenic axolotl tails were dissociated and incubated in L15 medium (Gibco) containing 30 U/mL papain (Sigma), 0.24 mg/mL cysteine, 40 μg/mL DNase I type IV, and 0.5 mg/mL trypsin inhibitor (Sigma) for 1 h at room temperature. Digestion was stopped by adding an equal amount of ovomucoid inhibitor [0.5 mg trypsin inhibitor (Sigma) and 0.5 mg/mL BSA (Sigma) in L15 medium (Gibco)]. The cells were triturated with a fire-polished Pasteur pipette, collected by centrifuga-

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tion for 5 min at 80 × g, resuspended in 1 mL neurosphere medium [DMEM/F12 (Gibco), 2% (vol/vol) B27 (Gibco), 20 ng/mL human recombinant FGF-2 (R & D Systems)], and plated at a density of 40,000 cells in 5 mL neurosphere medium. After 3 wk primary neurospheres either were plated onto dishes coated with poly-L-lysine (Sigma) and laminin (Sigma) for differentiation or were transplanted into the injured spinal cord of white (d/d) axolotls. Neurosphere Transplantation and Imaging. For dissection, axolotls were anesthetized in 0.03% (wt/wt) ethyl p-aminobenzoate (E-1501; Sigma) dissolved in water. An ∼2-mm-long portion of the spinal cord was removed from 4-cm-long white axolotls (d/d alleles; our colony) and enough EGFP+ neurosphere(s) to fill the gap were implanted (12). After 14 d of healing, tails were amputated and imaged weekly on an Olympus SZX 12 stereomicroscope using a cell-imaging system (Diagnostic Imaging Systems). After 2 mo axolotl tail samples were fixed in freshly made 4% (wt/wt) PFA at room temperature and then overnight at 4 °C. Then they were washed twice for 30 min each washing in 1× PBS and placed overnight in a solution of 30% (wt/wt) sucrose/1× PBS at 4 °C. For sectioning, the samples were embedded in TissueTek (Sakura). Clonal Neurosphere Formation and Implantation. The portions of tail spinal cord isolated from EGFP transgenic axolotls were dissociated as described above for the primary neurosphere cultures and were resuspended in neurosphere medium supplemented with 20 ng/mL FGF-2. To isolate the single EGFP+ cells from each other spatially, AggreWell 400 multiwell plates (STEMCELL Technologies) were used. Before cells were added, plates were centrifuged for 1 min at 1,000 × g in a plate centrifuge (Eppendorf Centrifuge 5804 with an A-2-DWP rotor) containing 600 μL neurosphere medium per well to remove all air bubbles. EGFP+ cells were mixed with white feeder cells at a ratio 1:5–1:10 and were distributed at a density of 600–1,000 EGFP+ cells per well. The plate was centrifuged at 100 × g for 2 min. The images of every single EGFP+ cell per microwell were taken on a Zeiss Axiovert inverted microscope with a Ph1 Plan Neofluar 10×/0.30 objective using the MetaMorph cell-imaging system (Molecular Devices). The cells were rechecked after 3–4 wk of culture to see if they formed clonal EGFP+ neurospheres. Individual clonal EGFP+ neurospheres were implanted in a white host spinal cord lesion (n = 28). After 1 wk of healing the tails were amputated close to the implant and were imaged every few days on an Olympus SZX 16 stereomicroscope using a cell-imaging system (Diagnostic Imaging Systems). After the process of regeneration had been completed, axolotl tail samples were fixed in freshly made 3.7% (wt/wt) formaldehyde in 1× MEM (Eagles’ minimum essential medium). After three washings for 30 min each in 1× PBS, they were placed overnight in a solution of 20% (wt/wt) sucrose/1× PBS at 4 °C. For cryosectioning, the samples were embedded in TissueTek (Sakura). Sox2 and HoxA9 A13 Antibody Preparation. A GST-fusion protein with amino acids 1–44 of the axolotl Sox2 was expressed in bacteria and purified by standard methods on Glutathione Sepharose (GSTrap; GE Healthcare). Rabbits were injected with 200 μg of purified protein in each of three boosts. Anti-serum was affinity purified using MBP-Sox2 fusion proteins conjugated to 1 mL of NHS-Sepharose resin (GE Healthcare). For HoxA9 antibody production, a HoxA9-GST fusion with amino acids RHYGIKPEPLPPGTRRGDCTTFDSSHTLSLSDYGSSPADKQSSEGAFPEAPAETEASGDKPAIDPNNPAANW was used to immunize rabbits. Antibody was affinity purified using a HoxA9-MBP fusion with amino acids PLPPGTRRGDCTTFDSSHTLSLSDYGSSPADKQSSEGAFPEAPAETEASGDKPA. For HoxA13, a GST-fusion protein with amino acids PTMVPTVGGPGEPRHEPLLPMEPYQPWALTNGWNGQVYCSKEQGQPPHLWKSSLPDVVSHPSDAN was used to immunize rabbits. Antibody was affinity purified using an MBP fusion protein to the same sequence. Specificity of antibody immunostaining was confirmed by comparing in situ hybridization pattern and immunostaining on developing limb buds. Immunohistochemistry. Neurosphere or tail sections were washed in PBS/0.1% (vol/vol) Tween and blocked in PBS/0.1% (vol/vol) Tween with 20% (vol/vol) goat serum or 1% (wt/wt) BSA. Sections were stained with primary antibodies against the following proteins: GFAP (1:500) (Chemicon); Musashi-1 (rat; 1:500) (gift of Hideyuki Okano, Keio University, Tokyo) (20); NeuN (1:1,000) (Chemicon), βIII-tubulin (mouse;1:500) (Chemicon); MBP (1:500) (GeneTex); PAX7 (Development Studies Hybridoma Bank); PAX6 (made in house); and MAP2 (mouse; 1:200) (Sigma). Appropriate secondary antibodies [goat antimouse Cy3, 1:100 (Chemicon); goat anti-mouse Cy5, 1:100 (Chemicon); and goat anti-rat Cy3, 1:100 (Dianova)] were applied and incubated for 1 h at room temperature. Nuclei were labeled with 1 μg/mL Hoechst for 15 min. Images were photographed with a Zeiss LSM UV laser-scanning confocal

PNAS | Published online July 24, 2012 | E2265

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spinal cord. Extensive studies of the hematopoietic stem cell system have shown that the hematopoietic system consists of hierarchies of stem cells, with only a very small fraction having ultimate, long-term ability to reconstitute the entire hematopoietic system (24, 25). Importantly, in irradiated animals stem cells without long-term reconstituting ability can act in a supportive role, allowing short-term survival so that rare cells can provide long-term reconstitution (25). Similarly, the spinal cord may have several types of stem cells, including a rare CNS/PNS stem cell. The nonclonal cultures, by providing a diversity of stem cells with different potencies, may robustly allow reconstitution of the DRG both via the presence of short-term reconstituting cells and the rare CNS/PNS stem cells, whereas the clonal cultures are not robust enough to allow efficient engraftment of the rare CNS/PNS stem cell. On the other hand, it also is possible that in the nonclonal neurosphere experiments a separate PNS-specific precursor that is closely associated with the spinal cord tissue (such as boundary cap cells) was included in the culture. The extensive reconstructive and neurogenic behavior of the axolotl system contrasts with mammalian neurosphere transplantation experiments that often led to astrocyte formation and limited neurogenesis (26–31). A future goal will be to determine whether these differences represent intrinsic differences in the stem cell populations or in the extrinsic cues present at the injury site. Histological characterization of the axolotl samples soon after implantation indicates that the cells integrate relatively rapidly with the endogenous radial glial cells to form a continuous cell layer. How this wound healing is achieved is an interesting question for future investigation.

microscope or Zeiss Apotome microscope and Zeiss Axiocam HR camera. All images were combined for figures using MetaView, Axiovision 4.6, and Photoshop CS (Adobe). ACKNOWLEDGMENTS. We thank Hideyuki Okano for providing antiMusashi antibodies; Jeremy Brockes for providing anti-HNK1; Verdon Taylor for introducing us to neurosphere-culturing techniques; Heino

Andreas for axolotl care; Regina Wegner for assistance with cell culture; and Hans H. Epperlein. This work was supported by funding from the Volkswagen Foundation Conditional Activation of Gene Expression; by Deutsche Forschungsgemeinschaft (DFG) Grants DFG-SFB655: Cells to Tissues, SPP1109 DFG TA-275/1-3, and DFG TA-274/3-1; and by institutional funds from the Max Planck Institute of Molecular Cell Biology and Geneticsand from the DFG Center for Regenerative Therapies Dresden.

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