Amputation-induced reactive oxygen species signaling is required for axolotl tail regeneration
Abstract
Background
Among vertebrates, salamanders are unparalleled in their ability to regenerate appendages throughput life. However, little is known about early signals that initiate regeneration in salamanders.
Results
Ambystoma mexicanum embryos were administered tail amputations to investigate the timing of reactive oxygen species (ROS) production and the requirement of ROS for regeneration. ROS detected by dihydroethidium increased within minutes of axolotl tail amputation and levels remained high for 24 hr. Pharmacological inhibition of ROS producing enzymes with diphenyleneiodonium chloride (DPI) and VAS2870 reduced ROS levels. Furthermore, DPI treatment reduced cellular proliferation and inhibited tail outgrowth.
Conclusions
The results show that ROS levels increase in response to injury and are required for tail regeneration. These findings suggest that ROS provide instructive, if not initiating cues, for salamander tail regeneration. Developmental Dynamics 248:189-196, 2019. © 2018 Wiley Periodicals, Inc.
Introduction
Urodele amphibians (i.e., salamanders) are unparalleled among vertebrates in their ability to regenerate appendages. Exactly how salamanders regenerate their limbs and tail largely remains a mystery. Many studies of salamander appendage regeneration have focused attention on the dynamics of progenitor cells that are recruited relatively late during the regeneration process (Mchedlishvili et al., 2007; Kragl et al., 2009). Earlier events that precede progenitor cell proliferation have received less attention. Several injury-associated cues affect the initiation of tissue regeneration in fish and frog tadpoles, including ion gradients, reactive oxygen species (ROS), and hypoxia (Tseng and Levin, 2012; Jopling et al., 2012; Love et al., 2013; Gauron et al., 2013; McLaughlin and Levin, 2018). Whether these or other cues initiate appendage regeneration in highly regenerative urodeles is unknown.
ROS are chemically reactive oxygen-containing molecules generated endogenously by several cellular components, including mitochondria, lyzosomes, and the plasma membrane (Di Meo et al., 2016). The reduction of molecular oxygen produces superoxide anion (O2-), a precursor of hydrogen peroxide (H2O2) and hydroxyl radicals (OH). ROS are well-known components of metazoan innate immune responses and, at moderate concentrations, serve others roles in tissue injury, wound healing, cellular signaling, and gene expression (Weidinger and Kozlov, 2015). A growing number of studies highlight ROS in the regeneration of organs that span vertebrate and invertebrate taxa, thus implicating ROS as a phylogenetically conserved mechanism that initiates regeneration programs (Niethammer et al., 2009; Gauron et al., 2013; Love et al., 2013; Piorette et al., 2015; Hameed et al., 2015; Zhang et al., 2016). These and other studies (Yoo et al., 2012; Lisse et al., 2016) show that ROS recruit regeneration-permissive lymphocytes, induce mitogen signaling pathways required for progenitor cell proliferation, and modulate cell fate. Thus, ROS are not only produced to mitigate pathogens at wound sites, additionally, they provide injury cues to initiate regeneration programs. Still, because relatively few models and research paradigms have been investigated to date, there is need to further examine ROS signaling during regeneration. Here, we report the requirement of ROS for appendage regeneration in the primary salamander model, Ambystoma mexicanum (axolotl).
Results and Discussion
Histology of Axolotl Embryo Tail Regeneration
Previously, we showed that developmental stage 42 (Bordzilovskaya et al., 1989) axolotl embryos are capable of regenerating 2 mm of amputated, distal tail tip tissue in approximately 7 days (Ponomareva et al., 2015). To further advance this model and provide context for interpreting chemical perturbation of the regeneration process, we used hematoxylin and eosin (H&E) staining to detail histological changes during tail regeneration (Fig. 1). The tail amputation procedure removed the symmetrical, rounded tail tip, leaving a blunt tail stump (Fig. 1A,A′). In approximately 40 min postamputation (mpa), the open wound was covered by a thin epithelium (Fig. 1B,B′). By 2 days postamputation (2 dpa), a few scattered erythrocytes and tissue debris were observed between the notochord and the wound epidermis (Fig. 1C,C′). Also at this time, mesenchymal tissue was observed between the wound epidermis and ependymal tube of the spinal cord, which had regenerated approximately 100 μm beyond the amputation plane. By 5 dpa, the spinal cord had lengthened considerably into the mesenchyme of the regenerating tail (Fig. 1D,D′). Ventral to the outgrowing spinal cord and distal to the notochord was a nodule of cartilaginous tissue observed at 7 dpa (Fig. 1E) that by 12 dpa was an elongated rod comprised of chondrocyte-like cells (Fig. 1F). Overall, histological changes observed during axolotl embryo tail regeneration approximated changes previously described for adult salamanders (Globus and Liversage, 1975; Iten and Bryant, 1976).
Association of ROS With Axolotl Embryo Tail Regeneration
ROS production during tail regeneration was measured by dihydroethidium (DHE), a dye that permeates plasma membranes to report super oxide anion production (Owusu-Ansah et al., 2008). Upon reaction with superoxide anions, DHE forms a red fluorescent product (ethidium) that intercalates with DNA and subsequently becomes retained by cells. After amputation of axolotl embryos, DHE staining revealed a marked increase of ROS production in midline, axial tissues (e.g., somites, spinal cord, notochord) of the tail (Fig. 2A). Elevated ROS levels remained high after wound closure and were sustained through 24 hr postamputation (hpa), but dropped to basal (unamputated) levels after that (Fig. 2B,C). This suggests that ROS production is associated with early wound healing events that precede cell proliferation and tail outgrowth.
NOX Activity is Required for Amputation Induced ROS Production
NAPDH oxidases (NOX) generate the majority of ROS (O2- and H2O2) (Bedard and Krause, 2007). These enzymes are phylogenetically conserved and expressed across many tissue types, and thus we used pharmacological inhibitors of NOX activity (diphenyleneiodonium chloride) [DPI] and VAS2870 to test the requirement of ROS for tail regeneration. DHE staining of embryos treated with 5 μM VAS2870 (Wingler et al., 2012) showed a significant reduction in ROS production at 6 hpa, however, embryos did not survive past 24 hpa. (Fig. 3). Lower, nonlethal doses of VAS2870 did not significantly lower ROS production or affect tail regeneration. Embryos treated with 4 μM DPI (Bedard and Krause, 2007) did not show a decrease in ROS production at 6 hpa, but levels were significantly lower at 24 hpa. Of interest, while a 0–24 hpa DPI treatment decreased ROS levels, embryos successfully completed tail regeneration. This suggested a requirement for NOX activity after 24 hpa. To test this hypothesis, embryos were treated for progressively longer periods of time, extending the treatment window by 24-hr intervals. Embryos that were treated for 0–2 days successfully completed tail regeneration while individuals that were treated for longer than 3 days died. Only individuals that were treated for 0–3 dpa showed significant inhibition of tail and spinal cord regeneration at 7 dpa (Fig. 4A,B). Because mesenchymal tissue did not form distal to the amputation plane in DPI-treated embryos, spinal cord regeneration was reduced and the underlying cartilaginous rod did not form (Fig. 4C). These results implicate NOX activity in amputation-induced ROS production and successful tail regeneration.
ROS as an Early Injury-Induced Signal and Its Role in Cellular Proliferation
ROS signaling is associated with the activation of cellular signaling pathways that regulate the proliferation of progenitor cells in planaria and frog tadpoles (Gauron et al., 2013; Love et al., 2013; Pirotte et al., 2015; Zhang et al., 2016). These findings prompted us to investigate the link between ROS production and cell proliferation during axolotl tail regeneration. The rate of distal tail outgrowth in the axolotl embryo model increases linearly between 12 and 72 hpa, and then after this time the rate increases more steeply (Ponomareva et al., 2015). Under the assumption that cell proliferation drives tail outgrowth, we quantified cell proliferation during the linear phase of tail outgrowth at 2 dpa. Embryos were treated with 4 μM DPI from 0–2 dpa and the number of mitotic, phospho-histone H3 (PH3) positive cells in the tail stump were quantified relative to untreated, control embryos (Fig. 5). Overall, relatively few mitotic cells were observed and their distribution was not localized within any particular tissue compartment. However, the number of PH3-postive cells was significantly higher in control embryos (Fig. 5A,B). Furthermore, we found that DPI treatment significantly reduced mitosis in the spinal cord and the epidermis but not in the mesenchyme (Fig. 5C,D). These results suggest that NOX activity affects cell mitosis in the spinal cord and the epidermis of the axolotl tail.
To further test the idea that NOX activity is required for cell proliferation, we tested DPI for an effect on cell cycle re-entry (S-phase). Embryos were treated continuously with 4 μM DPI, microinjected with 5-ethynyl-2'-deoxyuridine (EdU) at 2 dpa, and then tissue was collected at 3 dpa for quantification of Edu positive cells (Fig. 6A). EdU indices showed that cell proliferation was significantly reduced in DPI-treated embryos (Fig. 6B,C) relative to controls, which showed strong incorporation of EdU in regenerating spinal cord and tail fin (Fig. 6B, top panel). Although the area of regenerating tissue in control embryos was significantly greater than that of DPI-treated embryos at 3 dpa, EdU incorporation were still significantly greater in control embryos after normalizing for tissue area. While the regenerating tails of control embryos exhibited strong EdU incorporation in the regenerating spinal cord and the mesenchyme, DPI-treated embryos failed to regenerate spinal cord and had few EdU positive cells located within the tail fin (mesenchyme) (Fig. 6C). These results are similar to the data shown in (Fig. 5) which supports the idea that NOX activity and ROS signaling are required for cell cycle reentry and the proliferation response that drives regenerate outgrowth, especially in the spinal cord.
In the context of tissue damage or injury, ROS can modulate injury-induced gene expression responses and regulate proteins involved in wound signaling and regeneration (Enyedi and Niethammer, 2015; Neithammer, 2016; Lessi et al., 2016). Our results show that ROS signaling and NOX activity are required for axolotl spinal cord and tail regeneration. Although nothing is known about the regulatory targets of ROS signaling during axolotl tissue regeneration, studies of zebrafish and Xenopus suggest ROS may target essential mitogens, such as sonic hedgehog (SHH) and bone morphogenetic protein, which are required for proper cellular proliferation and blastema formation during axolotl tail regeneration (Schnapp et al., 2005; Stoick-Cooper et al., 2007). Recently, SHH was identified as a redox target during zebrafish fin regeneration in a study that examined ROS signaling during axon regeneration and cellular proliferation (Meda et al., 2016, 2017). Also, fgf20, one of the direct targets of Wnt/ β-catenin signaling, is down-regulated along with cell proliferation, when NOX-derived ROS production is inhibited during tadpole Xenopus tail regeneration (Love et al., 2013). Interestingly, Wnt signaling is essential for successful axolotl embryo tail regeneration and Fgf9 is one of the transcriptional targets that is down-regulated upon Wnt inhibition (Ponomareva et al., 2015). This implicates Wnt signaling as a potential candidate redox target of NOX activity and ROS production. Future studies will determine if these and other signaling pathways are regulated by ROS in the axolotl tail regeneration model.
Conclusions
Urodele amphibians are exceptional in regards to regenerating complex organs and tissues. Our results are consistent with findings from studies of planaria (Pirotte et al., 2015), zebrafish fin (Gauron et al., 2013), Xenopus tail (Love et al., 2013), and newt adult brain (Hameed et al., 2015); clearly ROS are required for successful tissue / appendage regeneration. We show that ROS are produced shortly after amputation in axial tissues and their production is sustained during the first 24 hpa. We also found that ROS signaling influences cellular proliferation during the early phase of tail regeneration. Our study is the first to show the requirement of NOX activity and ROS regulation for urodele appendage regeneration, and given similar findings in other metazoan taxa, suggest that ROS is a phylogenetically conserved mechanism of tissue regeneration.
Experimental Procedures
Ethical Procedures
The use of prefeeding stage axolotls does not require a protocol approved by the Institutional Animal Care and Use Committee at University of Kentucky; however, embryos used in this study were treated according to the same ethical standards that apply to feeding axolotls.
Regeneration Assay
Mexican axolotl embryos (RRID: AGSC_100E) were obtained from the Ambystoma Genetic Stock Center (RRID: SCR_006372) at University of Kentucky. Stage 42 (Bordzilovskaya et al., 1989) embryos were used for all experiments reported in this study. The embryos were manually hatched, administered benzocaine anesthesia (0.2 g in 10 ml ethanol / liter water) and photographed. Embryos with tail amputations were reared in artificial pond water (43.25 g NaCl, 0.625 g KCl, 1.25 g MgSO4, 2.5 g NaHCO3, and 1.25 g CaCl2 per 50 liters RO water). In administering amputations, the distal most 2 mm of tail tissue was removed with a sterile razor blade. Whole embryo images were captured using an Olympus SZX16 microscope with a 0.5× objective lens and DP400 camera. Embryos were reared at 18–19C° in 12-well microtiter plates, one embryo per well. The effect of 4 μM DPI on tail regeneration was assessed by measuring the distance between the distal tail tip and notochord, as the latter does not regenerate leaving a clear mark of the amputation plane. Experiments were performed three times; similar results were obtained and are combined for the analyses. Postamputation measurements were compared between chemically treated embryos (n = 15) and controls (n = 14) using Student's t-test, and P < 0.05 was considered statistically significant.
Histology and Immunohistochemistry
Embryos were killed in benzocaine and fixed in 4% paraformaldehyde (PFA) overnight at 4C°, washed three times with 1° phosphate buffered saline (PBS; pH = 7.4) and cryopreserved in 30% sucrose overnight at 4C°. Embryos were embedded in optimal cutting temperature compound and sectioned longitudinally at 10–15 μm. Tissue sections were mounted on slides and stained using hematoxylin and eosin (Fischer et al., 2008) or stored at -80C° for immunohistochemistry.
For immunohistochemistry, slides where rehydrated with 1× PBS, incubated with blocking buffer (1× PBS with 5% bovine serum albumin, 1% tween 20, and 0.3% Triton X-100; PBST) for 1 hr at room temperature. Slides were then incubated with mouse anti-PH3 (Ser10) antibody (1:1,000, EMD Millipore # 05-1336-S) overnight at 4C°. Slides were washed in 1× PBS and incubated with goat anti-mouse antibody (1:200, Thermo Fisher # A32723) for 1 hr at room temperature. Slides were incubated with DAPI (4′,6-diamidine-2-phenylidole-dihydrochloride; Thermo Fisher # 62248) in 1× PBS (1:25,000) for 20 min, washed in 1× PBS, and mounted with fluoroshield (Sigma-Aldrich # F6937).
In Vivo ROS Imaging
The detection of ROS during axolotl tail regeneration was performed by incubating embryos in 5 μM DHE in darkness for 2 hr before imaging. Embryos were then washed to remove excessive stain, anesthetized in 0.02% benzocaine, and imaged using an SZX16 Olympus microscope with 2× objective lens and DP400 camera. Fluorescence quantification at the tail tip was measured as described by Jensen (2013) and Ozkucur et al. (2010) using ImageJ (Schneider et al., 2012). Briefly, an exact region of interest was drawn around the distal tail for each embryo image and the gray pixel intensity was measured and background-subtracted. The measurements were then averaged for embryos (n = 6–12 embryos per time point) and results were analyzed by one-way analysis of variance followed by Tukey's post hoc analysis to compare all time points with preamputation levels. Fluorescence quantification of embryos treated with 5 μM VAS2870 (n = 15) and controls (n = 15), and embryos treated with 4 μM DPI (n = 15) and controls (n = 15) was performed as described above. All experiments were performed three times yielding similar results that were combined for statistical analysis.
The resulting data were analyzed using Student's t-test, and P < 0.05 was considered statistically significant.
Cellular Proliferation Assay
Cellular mitosis was measured using PH3 staining, where DPI-treated embryos and controls (n = 8 each) were fixed and processed as described above. Embryo tails were sectioned at 15–20 μM thickness and stained for PH3 as described above. For each embryo, three to eight sections were analyzed to identify anatomically matching sections between control and treated embryos. For each section, all cells staining positive for PH3 within the distal 0.5 mm2 of the amputation plane were counted as well as cells staining positive for DAPI (Franklin et al., 2017). A mitotic index was calculated by dividing the number of PH3 positive cells by the number of DAPI positive cells in a common area for each tissue section. Student's t-test was used to assess statistical significance P < 0.05 was considered significant. To further study the effect of DPI treatment on individual tissue types in the tail, we divided the number of PH3 positive cells within each of the following tissues (spinal cord, epidermis, and mesenchyme) by the number of cells found in the distal 0.5 mm2. Student's t-test was used to assess statistical significance, P < 0.05 was considered statistically significant.
To assay for cell cycle reentry, we used EdU Click-iT ®kit to detect DNA synthesis during S phase (Salic and Mitchison, 2008). Embryos at stage 42 were amputated and treated with either 4 μM DPI (n = 15) or 0.04% dimethyl sulfoxide (DMSO; n = 12). Embryos were microinjected with 0.5 μl of 8 mM EdU using a picospritzer at 2 dpa. Embryos were killed at 3 dpa and fixed in 4% PFA overnight. Embryos were washed three times in 1× PBS and dehydrated in a series of methanol concentrations (25%, 50%, 75%, 100%). Embryos were then rehydrated by running through the methanol concentrations backward and incubated in 1× trypsin for 30 min at room temperature. The embryos were then washed in distilled water three times and placed in acetone for 10 min at -20C° followed by washing in 1% PBST three times. EdU cocktail reaction was prepared per manufacturer kit with minor modification; 100 mM sodium ascorbate replaced Click-iT® EdU buffer additive at the same recommended volume per reaction. DAPI was used to counterstain nuclei. Embryos placed in EdU reaction solution were placed in the dark and gently rocked for 30 min followed by washing in 1× PBS before imaging.
Cell proliferation measurements were performed as described in (Franklin et al., 2017) with some modifications. We were interested in measuring cell proliferation within the regenerating tissue, distal to the amputation plane; these areas where used for quantification. All EdU labeled cells in regenerating tissue were counted as well as cells staining positive for DAPI, after normalizing for regenerate tissue area. Student's t-test was used to assess statistical significance and P < 0.05 was considered statistically significant.
