Abstract
In animal species that have the capability of regenerating tissues and limbs, cell proliferation is enhanced after wound healing and is essential for the reconstruction of injured tissue. Although the ability to induce cell proliferation is a common feature of such species, the molecular mechanisms that regulate the transition from wound healing to regenerative cell proliferation remain unclear. Here, we show that upon injury, InhibinβA and JunB cooperatively function for this transition during Xenopus tadpole tail regeneration. We found that the expression of inhibin subunit beta A (inhba) and junB proto-oncogene (junb) is induced by injury-activated TGF-β/Smad and MEK/ERK signaling in regenerating tails. Similarly to junb knockout (KO) tadpoles, inhba KO tadpoles show a delay in tail regeneration, and inhba/junb double KO (DKO) tadpoles exhibit severe impairment of tail regeneration compared with either inhba KO or junb KO tadpoles. Importantly, this impairment is associated with a significant reduction of cell proliferation in regenerating tissue. Moreover, JunB regulates tail regeneration via FGF signaling, while InhibinβA likely acts through different mechanisms. These results demonstrate that the cooperation of injury-induced InhibinβA and JunB is critical for regenerative cell proliferation, which is necessary for re-outgrowth of regenerating Xenopus tadpole tails.
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Introduction
Amphibians, such as Xenopus, can regenerate lost tissues after injury; this high capacity for regeneration has attracted attention for its potential applications in regenerative medicine1,2,3. Wound healing occurs as a cellular response to injury, and in regenerative animals, proliferation of regenerating cells is subsequently enhanced to restore lost tissue. It has been reported that inhibition of cell cycle progression after wound healing causes a severe delay in regeneration4,5, indicating that injury-induced cell proliferation is required for successful tissue regeneration. In neonatal mice, cardiomyocyte proliferation increases after cardiac apex resection; however, from one-week-old, the proliferation response does not occur in the mice even though the wound is closed, resulting in the failure of heart regeneration6. In adult mice, overexpression of cell cycle regulators after heart damage induces cardiomyocyte proliferation and improves heart function7. On the basis of these findings, it appears that injury-induced cell proliferation may be suppressed in non-regenerative animals. Therefore, the transition from wound healing to regenerative cell proliferation is a crucial process for tissue regeneration.
Xenopus tadpole tail regeneration is divided into two main phases: wound healing and proliferation. During the wound healing phase, Smad2/3 (signal transducers of TGF-β signaling) and ERK are known to be activated in response to injury, and these signaling factors play important roles in tissue regeneration4,8. After wound healing, tail regeneration proceeds to the proliferation phase, in which formation of the regeneration bud and regenerative outgrowth/cell proliferation occur. In the proliferation phase, morphogenetic signals, such as BMP, Wnt, and FGF, are critical for regenerative cell proliferation; these signals are also found to be important in other regenerative species3,9,10,11,12,13. In Xenopus tail regeneration, the genetic perturbation of BMP signaling reduces the number of proliferating cells in the regenerating tissue, and the activation of Wnt signaling promotes regenerative cell proliferation9,10. FGF acts downstream of BMP and Wnt signaling9, and it has been reported that FGF and BMP signaling are involved in cell proliferation during Xenopus limb regeneration14. Although regulatory factors that function in the wound healing and proliferation phases have been identified, the molecular mechanisms regulating the transition from wound healing to regenerative cell proliferation are not yet well understood.
We have previously shown that expression of junB proto-oncogene (junb) is induced by injury-activated TGF-β/Smad signaling during the wound healing phase and that JunB positively regulates cell proliferation in Xenopus tropicalis tail regeneration15. However, regenerative cell proliferation is not completely suppressed in junb KO tadpoles, suggesting that additional regulators promote cell proliferation in cooperation with JunB. In this study, we found that expression of both inhibin subunit beta A (inhba) and junb is induced through TGF-β/Smad and MEK/ERK signaling after injury. In addition, CRISPR/Cas9-mediated KO experiments revealed the cooperation of InhibinβA and JunB for regenerative outgrowth/cell proliferation. Interestingly, JunB regulates tail regeneration via FGF signaling, whereas the modulation of outgrowth/cell proliferation by InhibinβA is likely mediated by other mechanisms, acting either in parallel with or downstream of FGF signaling. In summary, the cooperative action of InhibinβA and JunB guides the transition from wound healing to regenerative cell proliferation in Xenopus tail regeneration. These findings shed light on the mechanisms underlying injury-induced cell proliferation in regenerative animals.
Results
The expression of inhba and junb is induced by injury-activated signaling
Although injury-induced JunB is important for regenerative outgrowth/cell proliferation in the Xenopus tadpole tail, a CRISPR/Cas9-mediated junb KO in F0 or compound heterozygous tadpoles is not sufficient to block tail regeneration15. This prompted us to search for regulators that cooperate with JunB for regenerative outgrowth/cell proliferation. Inhibition of TGF-β signaling in the regenerating tail after wound healing impairs cell proliferation in the regeneration bud4; thus, in addition to a TGF-β/Activin family ligand responsible for the wound healing phase, other TGF-β/Activin family ligands possibly work together with JunB to promote regeneration during the proliferation phase. To identify candidate TGF-β/Activin family ligands that cooperate with JunB for regenerative cell proliferation, we examined the levels of expression of ligands (Tgfβs, Inhibins, Nodals, Gdfs, and Myostatins) during X. tropicalis tadpole tail regeneration using published RNA-seq data16 (Supplementary Figure 1). Three genes belonging to the TGF-β/Activin family, namely inhba, tgfb1, and tgfb2, were identified as being highly expressed during the wound healing phase [~ 6 h post-amputation (hpa)] as is junb. However, only expression of inhba is upregulated after injury (Supplementary Figure 2). Since Smad2/3 and ERK are activated immediately after tail amputation4,8, we analyzed whether the injury-mediated increase in expression of inhba and junb is regulated by TGF-β/Smad and MEK/ERK signaling. As shown in Fig. 1, expression of inhba and junb during wound healing was downregulated by both SB-505124 (SB, TGF-β receptor inhibitor) and PD0325901 (PD, MEK/ERK inhibitor). These results suggest that, in addition to JunB, InhibinβA may also function downstream of injury-activated signaling to promote cell proliferation in Xenopus tail regeneration.
The expression of inhba and junb is regulated by injury-activated signaling during wound healing. qRT-PCR analysis of inhba (a) and junb (b) expression in DMSO (control), SB-505124 (SB), and PD0325901 (PD)-treated tails. The regenerating tails were isolated at 1 and 2 hpa for the expression of junb and inhba, respectively. The data were normalized against expression of rps18, and then by the value of DMSO. *P < 0.05, **P < 0.01.
As injury-activated TGF-β/Smad and MEK/ERK signaling are important for the induction of inhba and junb expression, we investigated the relationship between the activation of Smad2/3 and ERK after tail amputation. In contrast to tgfb1 KO tadpoles17, phosphorylated Smad2/3 (pSmad2/3) expression was not downregulated by PD treatment at 2 hpa (Supplementary Figure 3). It has been shown previously that ERK phosphorylation does not require TGF-β/Smad signaling during wound healing after amputation of the Xenopus tail8. Taken together, these observations suggest that activation of TGF-β/Smad signaling and of MEK/ERK signaling are unlikely to be dependent on each other at the beginning of wound healing but both are important for the coordinated expression of inhba and junb.
inhba is expressed in the regeneration bud of the tail
We examined the spatiotemporal expression pattern of inhba during Xenopus tail regeneration using whole-mount in situ hybridization (Fig. 2). A low level of inhba expression was observed in the tip of the uncut tail. After amputation, inhba expression was not detected until 1 hpa; clear inhba expression was observed near the amputation site at 2–4 hpa in the wound healing phase. From 6 hpa, inhba transcripts increased in the regenerating tail core tissue at the tip of the amputated tail stump. In the proliferation phase, inhba expression was detected in the regeneration bud and the posterior region of regenerating tail at 24 and 36–72 hpa, respectively. Sectioning of tadpoles after in situ hybridization showed that inhba was locally expressed in the regenerating mesenchyme, spinal cord, notochord, and epidermis (Supplementary Figure 4); this pattern of expression in regenerating tissue overlapped with that of junb. These data indicate that inhba is induced by tail amputation and is expressed during regeneration in the Xenopus tadpole tail.
Spatiotemporal expression pattern of inhba during Xenopus tail regeneration. Whole-mount in situ hybridization of inhba in uncut tails and in regenerating tails at 0, 0.25, 0.5, 1, 2, 4, 6, 9, 12, 24, 36, 48 and 72 hpa. Negative control using a sense probe shows no signals. Blue/purple signals indicate the expression of inhba. Black arrowheads indicate amputation sites. Scale bar, 200 μm.
InhibinβA is required for regenerative outgrowth/cell proliferation in cooperation with JunB
Since inhba was expressed in response to injury-activated signaling during tail regeneration, we investigated whether InhibinβA is essential for tail regeneration. We generated F0 inhba KO tadpoles using the CRISPR/Cas9 system: two different sgRNAs (sg 1 and sg 2) were designed using the genomic sequence corresponding to the pro-domain of InhibinβA; each sgRNA was injected into fertilized eggs (Fig. 3a). Both sgRNAs induced mutations in the inhba locus at moderate frequencies as determined by a T7 endonuclease I (T7E1) assay (Supplementary Figure 5). At 72 hpa, inhba KO tadpoles injected with each sgRNA showed a delay in tail regeneration, and the lengths of regenerating tails in inhba KO tadpoles were reduced compared to control tadpoles (tyrosinase (tyr) KO) (Fig. 3b and c). In addition, when classified according to the lengths of regenerating tails, a high proportion of inhba KO tadpoles showed delayed tail regeneration phenotypes (sg 1, 64.9%; sg 2, 74.0%) (Fig. 3d). The similarity of phenotypes of tadpoles injected with either sg 1 or sg 2 suggests that the retardation of tail regeneration in inhba KO tadpoles resulted from the specific inhibition of InhibinβA function. To increase the mutation efficiency, we injected a combination of sg 1 and sg 2 into fertilized eggs and found that 91.6% of alleles contained out-of-frame mutations with stop codons around the sgRNA target sites; 6.7% of alleles had in-frame mutations. In total, 98.3% of alleles had mutations in the inhba locus (Supplementary Figure 6). Moreover, the combination of sgRNAs increased the proportion of tadpoles displaying delayed tail regeneration (sg 1 + sg 2, 83.6%) compared with either sg 1 or sg 2 alone (Fig. 3d). Therefore, we used the combination of sg 1 and sg 2 in subsequent experiments. The delay in tail regeneration in inhba KO and junb KO tadpoles was specific as there was no significant delay in developing tadpole tails before tail amputation at stage 41/42; the delay in tail regeneration was observed even at 10 days post-amputation (dpa) (Supplementary Figure 7). In addition, the survival rate of these KO tadpoles was similar to that of the tyr KO control (Supplementary Figure 8). Finally, to further confirm the specificity of the phenotype observed in inhba KO, a rescue experiment was performed by overexpressing InhibinβA (Supplementary Figures 9 and 10). Overexpression of InhibinβA in inhba KO tadpoles partially but significantly rescued the shortened regenerating tail length and markedly reduced the proportion of tadpoles showing delayed tail regeneration. Taken together, we conclude that InhibinβA is required for regenerative outgrowth of the Xenopus tadpole tail.
InhibinβA is required for tail regeneration. (a) Schematic drawing of sgRNA target sites (sg 1 and sg 2) in the inhba locus. Grey boxes, untranslated regions; orange boxes, coding regions; arrows, single-guide RNA target sites; bar, intron region. (b) Representative phenotypes of tyr KO (control), inhba KO sg 1, inhba KO sg 2, and inhba KO sg 1 + sg 2 tadpoles at 72 hpa. (c) The lengths of regenerating tails in KO tadpoles at 72 hpa. (d) Summary of phenotypes in KO tadpoles at 72 hpa. On the basis of the lengths of regenerating tails at 72 hpa, tadpoles were classified into three phenotypic groups (normal regeneration, weakly delayed regeneration, or severely delayed regeneration). Black arrowheads indicate amputation sites. Scale bar, 200 μm. NS, not significant; ***P < 0.001.
Next, we examined whether InhibinβA and JunB cooperate for regenerative outgrowth/cell proliferation. Double KO (DKO) of inhba and junb was performed by combining two sets of sgRNAs (two each for inhba and junb). In the DKO tadpoles, the mutation efficiencies at inhba and junb loci were comparable to those observed in the respective single KO tadpoles based on the T7E1 assay. As shown in Fig. 4a, b, the lengths of regenerating tails were shorter in inhba/junb DKO tadpoles than in inhba KO or junb KO tadpoles. Furthermore, whole-mount immunostaining of phosphorylated Histone H3 (pH3) in regenerating cells at 36 hpa revealed that the number of proliferating cells was significantly lower in inhba/junb DKO tadpoles than in inhba KO or junb KO tadpoles (Fig. 4c, d). Notably, there were no significant differences in the rates of cell proliferation in the proximal (headward) tail region of amputation sites among experimental groups (Supplementary Figure 11; see also Fig. 5e, f described below), suggesting that InhibinβA and JunB function in regenerating tissue to regulate regenerative cell proliferation. These findings provide strong evidence that InhibinβA and JunB cooperate for regenerative outgrowth/cell proliferation in the Xenopus tadpole tail.
InhibinβA and JunB cooperate for regenerative outgrowth/cell proliferation. (a) Representative phenotypes of tyr KO (control), inhba KO, junb KO, and inhba/junb DKO tadpoles at 72 hpa. (b) The lengths of regenerating tails in KO and DKO tadpoles at 72 hpa. (c) Representative immunofluorescent images of pH3 staining (white dots) in KO and DKO tadpoles at 36 hpa. Whole-mount immunostaining was performed with the pH3 antibody; immunostaining without the pH3 antibody was used as a negative control (no 1st Ab). (d) Relative number of proliferating cells at 36 hpa. The number of pH3-positive cells was divided by the corresponding area. All values were normalized against the value of tyr KO. Black and white arrowheads indicate amputation sites. Scale bars, 200 μm. *P < 0.05, **P < 0.01, ***P < 0.001.
JunB regulates regenerative outgrowth/cell proliferation via FGF signaling. (a) Relative expression of fgf20 and tbxt in inhba KO and junb KO tadpoles at 36 hpa. The data were normalized against the expression of rps18, and then by the value of tyr KO. (b) Representative phenotypes of tyr KO (control), junb KO, junb KO + junb mRNA, and junb KO + fgf20 mRNA tadpoles at 72 hpa. (c) The lengths of regenerating tails in KO tadpoles at 72 hpa. (d) Summary of phenotypes in KO tadpoles at 72 hpa. On the basis of the lengths of regenerating tails at 72 hpa, tadpoles were classified into three phenotypic groups (normal regeneration, weakly delayed regeneration, or severely delayed regeneration). (e) Representative immunofluorescent images of pH3 staining (white dots) in tyr KO (control), junb KO, junb KO + junb mRNA, and junb KO + fgf20 mRNA tadpoles at 36 hpa. (f) Relative number of proliferating cells at 36 hpa. The number of pH3-positive cells was divided by the corresponding area. All values were normalized against the value of tyr KO. Black and white arrowheads indicate amputation sites. Scale bars, 200 μm. NS, not significant; *P < 0.05, **P < 0.01, ***P < 0.001.
JunB regulates regenerative outgrowth/cell proliferation through FGF signaling
To elucidate the molecular mechanisms by which InhibinβA and JunB initiate regeneration processes, we searched for candidate downstream factors of InhibinβA and JunB during Xenopus tail regeneration. Inhibition of FGF signaling is known to affect Xenopus tail regeneration9. The FGF ligand fgf20 is highly upregulated after wound healing, and morpholino-mediated knockdown of Fgf20 causes a delay in tail regeneration18. Moreover, regenerative outgrowth/cell proliferation of the zebrafish caudal fin is inhibited in fgf20a mutants19. We therefore examined whether InhibinβA and JunB regulate tail regeneration through FGF signaling. First, expression of fgf20 and tbxt (also known as brachyury), a downstream effector of FGF signaling, were analyzed in regenerating cells of inhba KO and junb KO tadpoles at 36 hpa. A quantitative reverse transcription-PCR (qRT-PCR) analysis showed that fgf20 and tbxt transcripts were downregulated to a significant extent in junb KO tadpoles (Fig. 5a), suggesting that JunB, but not InhibinβA, regulates the expression of fgf20 and tbxt at the time when regenerating cells were actively proliferating. A rescue experiment was performed to determine whether JunB facilitates tail regeneration through FGF signaling. We overexpressed Fgf20 or JunB in junb KO tadpoles and measured the lengths of regenerating tails in the tadpoles. Our results indicated that Fgf20 partially but significantly rescued the delay in tail regeneration of junb KO tadpoles to the same extent as JunB (Fig. 5b, c; Supplementary Figure 10). Moreover, when classified according to the lengths of regenerating tails, the proportion of junb KO tadpoles with severely delayed tail regeneration was greatly reduced by Fgf20 overexpression (Fig. 5d). Next, we analyzed cell proliferation in these tadpoles at 36 hpa. As shown in Fig. 5e, f, the reduction in proliferating cells in junb KO tadpoles was rescued by overexpression of Fgf20 or JunB. Together with the finding of downregulated expression of fgf20 and tbxt in junb KO tadpoles (described above), these data support the conclusion that JunB acts, at least in part, through FGF signaling to promote regenerative outgrowth/cell proliferation of the Xenopus tadpole tail.
FGF signaling in the proliferation phase is important for regenerative outgrowth/cell proliferation
FGF signaling is well known to be essential for tissue/organ regeneration in regenerative animals9,12,14,18,19,20. Although inhibition of FGF signaling causes a delay in Xenopus tail regeneration9, the role of FGF signaling in regenerative cell proliferation is still unclear. In addition, the expression of fgf20, which functions downstream of injury-induced JunB, increases as outgrowth/cell proliferation progresses in the regenerating Xenopus tail18, and inhibition of FGF signaling does not affect wound healing9. Therefore, we speculated that FGF signaling in the proliferation phase might be required for outgrowth/cell proliferation in Xenopus tail regeneration. To suppress FGF signaling specifically in the proliferation phase, tail-amputated tadpoles were treated with SU5402 (SU, FGF receptor inhibitor) from 24 hpa (the time when regenerative outgrowth begins) and the lengths of regenerating tails were measured at 72 hpa (Fig. 6a). Consistent with a previous report9, inhibition of FGF signaling in the proliferation phase caused a delay in tail regeneration compared with DMSO-treated tadpoles (Fig. 6b, c). We then analyzed cell proliferation in SU-treated tadpoles and found that the number of proliferating cells in the regenerating tail was considerably reduced by SU (Fig. 6d). Thus, our results demonstrate that FGF signaling in the proliferation phase is essential for regenerative outgrowth/cell proliferation of the Xenopus tadpole tail. Given the fact that JunB, but not InhibinβA, regulates tail regeneration via Fgf20 (Fig. 5), we conclude that FGF signaling downstream of JunB functions in parallel with InhibinβA signaling in the proliferation phase (Fig. 7; see Discussion).
FGF signaling is required for regenerative outgrowth/cell proliferation. (a) Schematic of the experimental plan. Tadpoles were incubated in water containing 0.0375% DMSO (control) or 15 μM SU5402 (SU) from 24 hpa and analyzed at 72 hpa for regenerating tail length and proliferating cells. (b) Representative phenotypes of DMSO and SU-treated tadpoles at 72 hpa. (c) The lengths of regenerating tails at 72 hpa. (d) Relative number of proliferating cells at 72 hpa. The number of pH3-positive cells was divided by the corresponding area. In (b) and (c), the photograph and length of regenerating tails were obtained after whole-mount immunostaining, which involves bleaching and methanol treatments. Due to these treatments, the measured regenerating tail length is relatively shorter than in other experiments. All values were normalized against the value of DMSO-treated tadpoles. Black arrowheads indicate amputation sites. Scale bar, 200 μm. **P < 0.01, ***P < 0.001.
A model for the transition from wound healing to regenerative cell proliferation mediated by InhibinβA and JunB. Upon tail amputation, Tgfβ1/TGF-β receptor (TGF-βR) and MEK independently phosphorylate/activate Smad2/3 (pSmad2/3) and ERK (pERK), respectively. These injury-activated signals coordinately induce the expression of inhba and junb during the wound healing phase. After wound healing, InhibinβA and JunB cooperate to initiate outgrowth and cell proliferation. Mechanistically, JunB promotes tail regeneration, at least in part, via Fgf20/FGF receptor (FGFR) signaling; InhibinβA/Activin receptor (ActR) signaling may act either in parallel with or downstream of Fgf20/FGFR signaling (see Discussion).
Discussion
Injury-induced cell proliferation is a shared feature in animal species that have the ability to regenerate tissues. However, it is still not clear what factors are induced in response to injury to promote cell proliferation during tissue regeneration. Here, we show that inhba and junb are induced through injury-activated TGF-β/Smad and MEK/ERK signaling during the wound healing phase. Moreover, our study suggests the cooperative function of InhibinβA and JunB in injury-induced cell proliferation and tail regeneration. We propose that InhibinβA and JunB are important factors that mediate the transition from wound healing to regenerative cell proliferation during Xenopus tail regeneration (Fig. 7). The significance and possible mechanisms of the cooperative regulation of cell proliferation by InhibinβA and JunB are discussed in detail below.
Upon injury, multiple cell signaling pathways (TGF-β/Smad, ERK, and ROS) are known to be activated in the wound healing phase of the regenerating Xenopus tail4,8,18. However, the molecular mechanisms by which these injury-activated signals control regenerative outgrowth/cell proliferation through the regulation of gene expression are not well understood. We found that at the beginning of wound healing (~ 2 hpa), the phosphorylation of Smad2/3 was not dependent on MEK/ERK signaling (Supplementary Figure 3). Conversely, a previous report showed that the phosphorylation of ERK is not affected by inhibition of TGF-β/Smad signaling8. These observations imply that TGF-β/Smad and MEK/ERK signaling are independently activated by injury and coordinately regulate the expression of inhba and junb during wound healing (Figs. 1 and 7). While the upstream factors that activate MEK/ERK have not been elucidated, Tgfβ1 is required for the injury-induced phosphorylation of Smad2/3 (Supplementary Figure 3)17. Interestingly, tgfb1 is already highly expressed in the uncut Xenopus tail4,17; after tail amputation, Tgfβ1 stored in the extracellular matrix is expected to induce expression of downstream target genes, including inhba and junb, via the activation of Smad2/3. We postulate that MEK/ERK signaling may help to refine the expression of inhba and junb induced by the Tgfβ1-Smad2/3 pathway. The injury-induced burst of inhba and junb expression is fast and takes place within a few hours (~ 6 hpa). Therefore, it is possible that the coordination of TGF-β/Smad and MEK/ERK signaling is essential for a sharp and sustained increase in inhba and junb expression after tail amputation, ultimately contributing to the successful recovery from injury and the regeneration of damaged tissue.
The induction of inhba expression by the Tgfβ1-Smad2/3 pathway indicates the amplification of TGF-β/Smad signaling during Xenopus tail regeneration (Fig. 7). This is consistent with the observation that expression of TGF-β family ligands (inhba and gdf11) is sequentially induced during tail regeneration4. As described above, the activation of Smad2/3 is not suppressed by inhibition of MEK/ERK signaling at the beginning of wound healing. In contrast, at the end of wound healing, the phosphorylation of Smad2/3 is reduced by MEK/ERK inhibitor8. We speculate that the reduction of activated Smad2/3 at the late time point of wound healing may be due to the disruption of the amplification system of TGF-β/Smad signaling by downregulation of inhba after MEK/ERK inhibition (Fig. 1a). Furthermore, the core components of TGF-β signaling (ligands, receptors, and their transcriptional target genes) were found to be co-expressed during axolotl limb regeneration21. Given that TGF-β signaling plays critical roles in multiple steps of regeneration processes4,22,23,24, it is tempting to suggest that the amplification of TGF-β/Smad signaling is conserved among animal species with a high regenerative capacity and is fundamental for maintaining TGF-β signaling throughout regeneration.
Unlike urodeles, regenerating axial tissues (spinal cord, notochord, and somite) in the Xenopus tail have been shown to originate from the respective differentiated tissues in the tail stump, and metaplasia of tissue progenitors is not observed25. In addition, dedifferentiation phenotypes are not found and regenerating myofibers are derived from skeletal muscle tissue stem cells (satellite cells). These observations indicate that tissue renewal mediated by the induction of cell proliferation in differentiated cells is a critical step of Xenopus tail regeneration. Therefore, while InhibinβA and JunB could regulate differentiation directly or indirectly at some stages of regeneration, it is plausible that the cooperative function of InhibinβA and JunB in regenerative cell proliferation is important for successful tissue recovery. It will be interesting to analyze the contribution of this cooperation in other animal species with different regenerative capacities and cellular plasticities, particularly in urodeles.
During spinal cord regeneration in the axolotl, cell proliferation is accelerated by the progression of the cell cycle from the G0/G1 to S phase26, indicating that the release from the G0/G1 phase is a crucial step in regenerative cell proliferation. Notably, it has been shown that inhibition of InhibinβA or JunB in cultured cell lines increases the proportion of cells in the G0/G1 phase27,28. Moreover, in the present study, impairment of regenerative cell proliferation in inhba/junb DKO was greater than in either inhba KO or junb KO (Fig. 4d). Therefore, in response to tail amputation, InhibinβA and JunB may work together in Xenopus to promote cell cycle re-entry by facilitating the release from the G0/G1 phase. It has been reported that both Smad2/3 (signal transducers of InhibinβA) and JunB are essential for the transcription of cell cycle-related genes29,30 and that Smad3 regulates target gene expression by binding to JunB31,32,33,34. These findings and our results suggest that the cooperative function of InhibinβA and JunB in regenerative cell proliferation might be mediated by a transcriptional complex composed of Smad2/3 and JunB during Xenopus tail regeneration. In this case, InhibinβA-activated Smad2/3 and JunB directly converge in the cell cycle regulatory system. In parallel with this potential mechanism, InhibinβA may also interact with downstream effectors of JunB in the regulation of cell proliferation. Intriguingly, we found that Fgf20 acted downstream of JunB in Xenopus tail regeneration (Fig. 5). In addition, Activin A, a homodimer of InhibinβA, requires FGF signaling in mesoderm formation during Xenopus embryogenesis and in the endodermal differentiation of mouse embryogenic stem cells35,36,37. As the expression of fgf20 and its target gene tbxt was not reduced in inhba KO tadpoles when regenerating cells were actively proliferating (Fig. 5a), it is possible that InhibinβA functions with FGF signaling downstream of tbxt expression to regulate tail regeneration. Tbxt is known to associate with Smad2/3, and the genomic binding sites of Tbxt overlap with those of Smad2/3 in differentiating human embryonic stem cells38. An alternative but not mutually exclusive possibility is that InhibinβA affects other signaling pathways involved in regeneration processes. During regeneration of the zebrafish fin and Xenopus tadpole tail, Activin/TGF-β signaling regulates the expression of the BMP target gene msx14,24. Since BMP signaling is required for Xenopus tail regeneration10, InhibinβA might promote tail regeneration in cooperation with JunB by modulating the BMP pathway. Overall, the collaborative action of InhibinβA and JunB in regenerative outgrowth/cell proliferation may be mechanistically important for the robust patterning of regenerating tissue; this possibility will be investigated in future studies.
In conclusion, we showed that inhba and junb are coordinately induced upon tail amputation through TGF-β/Smad and MEK/ERK signaling, and that injury-induced InhibinβA and JunB cooperate to drive regenerative outgrowth/cell proliferation in the Xenopus tadpole tail. These findings provide valuable insights into the molecular mechanisms that mediate the transition from wound healing to regenerative cell proliferation during tissue regeneration.
Methods
Animals, microinjection, and regeneration assay
All experiments were conducted in accordance with ARRIVE guidelines and with the Fundamental Guidelines for Proper Conduct of Animal Experiment and Related Activities in Academic Research Institutions under the jurisdiction of the Ministry of Education, Culture, Sports, Science and Technology of Japan, and were approved by the Hiroshima University Animal Research Committee (Permit Number: G19-7-2). X. tropicalis embryos were cultured in 0.1X Marc’s Modified Ringer solution (MMR) at 26 °C. Tadpoles at stage 41/42 were anesthetized in 0.05% MS-222/0.01X MMR and subjected to tail amputation at the mid-point (50%) of tail length using a surgical knife. The removal of as much as 75% of tail length does not affect tail regeneration or the survival of tadpoles39. The amputated tadpoles were maintained until 72 hpa or 10 dpa in tap water. For inhibitor experiments, tadpoles were treated with 25 μM SB-505124 (Cayman Chemical) or 0.25 μM PD0325901 (Cayman Chemical) from 1 h before tail amputation, or with 15 μM SU5402 (Sigma) from 24 hpa.
Cloning of X. tropicalis inhba and fgf20
Full-length cDNAs for inhba (GenBank Accession number XM_002933406.3) and fgf20 (GenBank Accession number NM_001143927.1) were PCR-amplified using the following primers: inhba, forward 5′-CGG GAT CCC ACC TGG TGA CAG GAT GC-3′ and reverse 5′-GCT CTA GAA ATT GCT GCA GGC TGG TAA C-3′; fgf20, forward 5′-CGG GAT CCC TTT TGG GGA TTT TGG GAC T-3′ and reverse 5′-AAG GCC TGC ACT GGG TTT GGT TTG TCT-3′. The PCR products from inhba and fgf20 were cloned into the BamHI/XbaI and BamHI/StuI sites of pDH105 (pDH105-inhba and pDH105-fgf20), respectively.
CRISPR/Cas9-mediated KO, rescue experiment, and genotyping
CRISPR/Cas9-mediated mutagenesis was carried out as previously described15,40. For the KO experiment, 1000 pg of sgRNA was used. In the DKO experiment, 500 pg each of the two different sgRNAs for inhba and junb were used; the total amount of sgRNAs was 2000 pg. The injection solution was prepared by mixing sgRNAs and 1 ng of Cas9 protein (Integrated DNA Technologies) per 1000 pg of sgRNAs. For the rescue experiment, the capped junb, inhba, and fgf20 mRNAs were synthesized from pDH105-junb41, pDH105-inhba, and pDH105-fgf20, respectively, using the SP6 transcription kit (Invitrogen). mRNA was mixed with sgRNAs and Cas9 protein, and the mixture was injected into the fertilized eggs. The sequences of sgRNAs for tyr, junb, and tgfb1 were described in previous studies15,17,40, and primers for generating inhba sgRNAs were as follows: sg 1, 5′-ATT TAG GTG ACA CTA TAG GCC CCA ACT CCA GGA TCT GGT TTT AGA GCT AGA AAT AGC AAG-3′; sg 2, 5′-ATT TAG GTG ACA CTA TAG GCC ATG TCA CTC TGA GAA CGT TTT AGA GCT AGA AAT AGC AAG-3′. Genomic DNA was extracted from inhba KO tadpoles at 72 hpa and purified by the GenElute mammalian genomic DNA miniprep kit (Sigma). PCR amplification of the genomic region of inhba was performed using Q5 High Fidelity DNA polymerase (New England Biolabs). The PCR products were treated with T4 polynucleotide kinase (New England Biolabs) and cloned into the pUC57 vector, which was digested by EcoRV and dephosphorylated with Quick CIP (New England Biolabs). To determine the mutation types, single colonies were subjected to Sanger sequencing. T7E1 (New England Biolabs) assay was performed as previously described15. The following primers were used for PCR and sequencing: inhba, forward 5′-ACA GCC ACA AAT ACC CAC AG-3′ and reverse 5′-AAG GAG CCA GTG AAG CTT TG-3′.
RNA isolation and qRT-PCR
RNA isolation was performed as previously described15. In brief, regenerating tails were surgically isolated from 15 to 30 Xenopus tadpoles and lysed in a solution containing proteinase K. RNA was purified by phenol/chloroform extraction and ethanol precipitation. cDNA was synthesized using the PrimeScript RT reagent kit (Takara), and qRT-PCR was performed using THUNDERBIRD Next SYBR qPCR Mix (TOYOBO) and CFX Connect Real-Time System (BIO-RAD). Each qRT-PCR experiment was performed in triplicate. To detect endogenous mRNAs, the following primers were used for qRT-PCR: inhba, forward 5′-CAT GTG ATC AAT GCC AGG AG-3′ and reverse 5′-TCT TCC TCA ACA CCA GAT GC-3′; fgf20, forward 5′-TCG CAT CTC CAG GGA ATC-3′ and reverse 5′-GAT GAA TTC AAG GAT ACC AAA GC-3′; tbxt, forward 5′-CAA TTC CCC AAA CAA TTT AGC-3′ and reverse 5′-TGA CTT GAG GTA GGA GGT GTT C-3′; the primer sequences of junb and rps18 were described in the previous study15. To detect injected mRNAs, the following primers were used: inhba, forward 5′-AAG TCA GAG TTA TCT GGG TTT GG-3′ and reverse 5′-GCA TTC TAG TTG TGG TTT GTC C-3′; fgf20, forward 5′-GCC AAA GAC AAA CCA AAC CCA-3′ and reverse 5′-TGG ATC TAC GTA ATA CGA CTC ACT-3′; junb, forward 5′-TTA CTG GGA AAG GGC ATC AG-3′ and reverse 5′-GCA TTC TAG TTG TGG TTT GTC C-3′.
Whole-mount in situ hybridization and whole-mount immunostaining
Whole-mount in situ hybridization and cryosectioning of hybridized tadpoles were performed as previously described42,43,44. Tadpoles were fixed in MEMFA solution for 2 h at room temperature and stored in 100% ethanol at − 20 °C. For anti-sense and sense probes, pDH105-inhba was linearized by BamHI and XbaI and subjected to in vitro transcription by T3 and SP6 RNA polymerases (Roche), respectively. For the whole-mount immunostaining, tadpoles were fixed in MEMFA solution for 30 min at room temperature and stored in 100% methanol at − 20 °C. Whole-mount immunostaining was carried out following our standard protocol for pH3 staining15 and with a minor modification for pSmad2/3 staining17. The following antibodies were used at 1:500 dilution: anti-pH3 antibody (Upstate Biotechnology), anti-pSmad2/3 antibody (Cell Signaling Technology), Alexa Fluor 488 goat anti-rabbit antibody (Molecular Probes, Thermo Fisher Scientific), and Alexa Fluor 488 goat anti-mouse antibody (Molecular Probes, Thermo Fisher Scientific).
Microscopy, quantification, and statistical analysis
Fluorescent images were obtained with Axio Zoom.V16 (ZEISS), and the fluorescence intensity was quantified with ZEN blue software (ZEISS). Images of cryosections were captured using an Axio Observer.Z1 (ZEISS). Quantification of pSmad2/3 staining was performed as described previously17. The length of the regenerating tail was measured with cellSens Standard imaging software (Olympus) or ImageJ software (National Institutes of Health, USA). On the basis of the lengths of regenerating tails at 72 hpa, tadpoles were classified as showing normal regeneration (more than 80% of the average length of tyr KO), weakly delayed regeneration (60–80% of the average length of tyr KO), or severely delayed regeneration (less than 60% of the average length of tyr KO). The number of pH3-positive cells was manually counted in the regenerating tail, excluding the fin. Statistical analysis was performed by Student’s t-test with or without Bonferroni correction, and error bars indicate the standard error of the mean.
Data availability
No dataset was generated in this study. The GenBank accession numbers of inhba and fgf20 genes isolated in this study are given in the Methods.
References
Kakebeen, A. D. & Wills, A. E. More than just a bandage: closing the gap between injury and appendage regeneration. Front. Physiol. 10, 81. https://doi.org/10.3389/fphys.2019.00081 (2019).
Phipps, L. S., Marshall, L., Dorey, K. & Amaya, E. Model systems for regeneration: Xenopus. Development 147, dev180844. https://doi.org/10.1242/dev.180844 (2020).
Bassat, E. & Tanaka, E. M. The cellular and signaling dynamics of salamander limb regeneration. Curr. Opin. Cell Biol. 73, 117–123. https://doi.org/10.1016/j.ceb.2021.07.010 (2021).
Ho, D. M. & Whitman, M. TGF-β signaling is required for multiple processes during Xenopus tail regeneration. Dev. Biol. 315, 203–216. https://doi.org/10.1016/j.ydbio.2007.12.031 (2008).
Wang, Y.-T. et al. Genetic reprogramming of positional memory in a regenerating appendage. Curr. Biol. 29, 4193–4207. https://doi.org/10.1016/j.cub.2019.10.038 (2019).
Porrello, E. R. et al. Transient regenerative potential of the neonatal mouse heart. Science 331, 1078–1080. https://doi.org/10.1126/science.1200708 (2011).
Mohamed, T. M. A. et al. Regulation of cell cycle to stimulate adult cardiomyocyte proliferation and cardiac regeneration. Cell 173, 104–116. https://doi.org/10.1016/j.cell.2018.02.014 (2018).
Sato, K., Umesono, Y. & Mochii, M. A transgenic reporter under control of an es1 promoter/enhancer marks wound epidermis and apical epithelial cap during tail regeneration in Xenopus laevis tadpole. Dev. Biol. 433, 404–415. https://doi.org/10.1016/j.ydbio.2017.08.012 (2018).
Lin, G. & Slack, J. M. W. Requirement for Wnt and FGF signaling in Xenopus tadpole tail regeneration. Dev. Biol. 316, 323–335. https://doi.org/10.1016/j.ydbio.2008.01.032 (2008).
Beck, C. W., Christen, B., Barker, D. & Slack, J. M. W. Temporal requirement for bone morphogenetic proteins in regeneration of the tail and limb of Xenopus tadpoles. Mech. Dev. 123, 674–688. https://doi.org/10.1016/j.mod.2006.07.001 (2006).
Wagner, I. et al. Serum proteases potentiate BMP-induced cell cycle re-entry of dedifferentiating muscle cells during newt limb regeneration. Dev. Cell 40, 608–617. https://doi.org/10.1016/j.devcel.2017.03.002 (2017).
Nacu, E., Gromberg, E., Oliveira, C. R., Drechsel, D. & Tanaka, E. M. FGF8 and SHH substitute for anterior-posterior tissue interactions to induce limb regeneration. Nature 533, 407–410. https://doi.org/10.1038/nature17972 (2016).
Stoick-Cooper, C. L., Moon, R. T. & Weidinger, G. Advances in signaling in vertebrate regeneration as a prelude to regenerative medicine. Genes Dev. 21, 1292–1315. https://doi.org/10.1101/gad.1540507 (2007).
Mitogawa, K., Makanae, A. & Satoh, A. Hyperinnervation improves Xenopus laevis limb regeneration. Dev. Biol. 433, 276–286. https://doi.org/10.1016/j.ydbio.2017.10.007 (2018).
Nakamura, M. et al. The AP-1 transcription factor JunB functions in Xenopus tail regeneration by positively regulating cell proliferation. Biochem. Biophys. Res. Commun. 522, 990–995. https://doi.org/10.1016/j.bbrc.2019.11.060 (2020).
Chang, J., Baker, J. & Wills, A. Transcriptional dynamics of tail regeneration in Xenopus tropicalis. Genesis 55, e23015. https://doi.org/10.1002/dvg.23015 (2017).
Nakamura, M. et al. TGF-β1 signaling is essential for tissue regeneration in the Xenopus tadpole tail. Biochem. Biophys. Res. Commun. 565, 91–96. https://doi.org/10.1016/j.bbrc.2021.05.082 (2021).
Love, N. R. et al. Amputation-induced reactive oxygen species are required for successful Xenopus tadpole tail regeneration. Nat. Cell Biol. 15, 222–228. https://doi.org/10.1038/ncb2659 (2013).
Whitehead, G. G., Makino, S., Lien, C.-L. & Keating, M. T. fgf20 is essential for initiating zebrafish fin regeneration. Science 310, 1957–1960. https://doi.org/10.1126/science.1117637 (2005).
Prudovsky, I. Cellular mechanisms of FGF-stimulated tissue repair. Cells 10, 1830. https://doi.org/10.3390/cells10071830 (2021).
Tsai, S. L., Baselga-Garriga, C. & Melton, D. A. Blastemal progenitors modulate immune signaling during early limb regeneration. Development 146, dev169128. https://doi.org/10.1242/dev.169128 (2019).
Chablais, F. & Jaźwińska, A. The regenerative capacity of the zebrafish heart is dependent on TGFβ signaling. Development 139, 1921–1930. https://doi.org/10.1242/dev.078543 (2012).
Lévesque, M. et al. Transforming growth factor: β signaling is essential for limb regeneration in axolotls. PLoS ONE 2, e1227. https://doi.org/10.1371/journal.pone.0001227 (2007).
Jaźwińska, A., Badakov, R. & Keating, M. T. Activin-βA signaling is required for zebrafish fin regeneration. Curr. Biol. 17, 1390–1395. https://doi.org/10.1016/j.cub.2007.07.019 (2007).
Gargioli, C. & Slack, J. M. W. Cell lineage tracing during Xenopus tail regeneration. Development 131, 2669–2679. https://doi.org/10.1242/dev.01155 (2004).
Costa, E. C., Otsuki, L., Albors, A. R., Tanaka, E. M. & Chara, O. Spatiotemporal control of cell cycle acceleration during axolotl spinal cord regeneration. eLife 10, e55665. https://doi.org/10.7554/eLife.55665 (2021).
Zhu, J., Liu, F., Wu, Q. & Liu, X. Activin A regulates proliferation, invasion and migration in osteosarcoma cells. Mol. Med. Rep. 11, 4501–4507. https://doi.org/10.3892/mmr.2015.3284 (2015).
Zhang, J. et al. The c-Jun and JunB transcription factors facilitate the transit of classical Hodgkin lymphoma tumour cells through G1. Sci. Rep. 8, 16019. https://doi.org/10.1038/s41598-018-34199-9 (2018).
Park, Y. et al. Induction of cyclin D2 in rat granulosa cells requires FSH-dependent relief from FOXO1 repression coupled with positive signals from Smad. J. Biol. Chem. 280, 9135–9148. https://doi.org/10.1074/jbc.M409486200 (2005).
Andrecht, S., Kolbus, A., Hartenstein, B., Angel, P. & Schorpp-Kistner, M. Cell cycle promoting activity of JunB through cyclin A activation. J. Biol. Chem. 277, 35961–35968. https://doi.org/10.1074/jbc.M202847200 (2002).
Liberati, N. T. et al. Smads bind directly to the Jun family of AP-1 transcription factors. Proc. Natl. Acad. Sci. USA 96, 4844–4849. https://doi.org/10.1073/pnas.96.9.4844 (1999).
Selvamurugan, N., Kwok, S. & Partridge, N. C. Smad3 interacts with JunB and Cbfa1/Runx2 for transforming growth factor-β1-stimulated collagenase-3 expression in human breast cancer cells. J. Biol. Chem. 279, 27764–27773. https://doi.org/10.1074/jbc.M312870200 (2004).
Sundqvist, A. et al. Specific interactions between Smad proteins and AP-1 components determine TGFβ-induced breast cancer cell invasion. Oncogene 32, 3606–3615. https://doi.org/10.1038/onc.2012.370 (2013).
Sundqvist, A. et al. JUNB governs a feed-forward network of TGFβ signaling that aggravates breast cancer invasion. Nucl. Acid. Res. 46, 1180–1195. https://doi.org/10.1093/nar/gkx1190 (2018).
Cornell, R. A. & Kimelman, D. Activin-mediated mesoderm induction requires FGF. Development 120, 453–462. https://doi.org/10.1242/dev.120.2.453 (1994).
Hansson, M. et al. A late requirement for Wnt and FGF signaling during activin-induced formation of foregut endoderm from mouse embryonic stem cells. Dev. Biol. 330, 286–304. https://doi.org/10.1016/j.ydbio.2009.03.026 (2009).
Funa, N. S., Saldeen, J., Åkerblom, B. & Welsh, M. Interdependent fibroblast growth factor and activin A signaling promotes the expression of endodermal genes in differentiating mouse embryonic stem cells expressing Src Homology 2-domain inactive Shb. Differentiation 76, 443–453. https://doi.org/10.1111/j.1432-0436.2007.00249.x (2008).
Faial, T. et al. Brachyury and SMAD signalling collaboratively orchestrate distinct mesoderm and endoderm gene regulatory networks in differentiating human embryonic stem cells. Development 142, 2121–2135. https://doi.org/10.1242/dev.117838 (2015).
Beck, C. W., Belmonte, J. C. I. & Christen, B. Beyond early development: Xenopus as an emerging model for the study of regenerative mechanisms. Dev. Dyn. 238, 1226–1248. https://doi.org/10.1002/dvdy.21890 (2009).
Blitz, I. L., Biesinger, J., Xie, X. & Cho, K. W. Y. Biallelic genome modification in F0 Xenopus tropicalis embryos using the CRISPR/Cas system. Genesis 51, 827–834. https://doi.org/10.1002/dvg.22719 (2013).
Yoshida, H., Okada, M., Takebayashi-Suzuki, K., Ueno, N. & Suzuki, A. Involvement of JunB proto-oncogene in tail formation during early Xenopus embryogenesis. Zool. Sci. 33, 282–289. https://doi.org/10.2108/zs150136 (2016).
Takebayashi-Suzuki, K., Kitayama, A., Terasaka-Iioka, C., Ueno, N. & Suzuki, A. The forkhead transcription factor FoxB1 regulates the dorsal-ventral and anterior-posterior patterning of the ectoderm during early Xenopus embryogenesis. Dev. Biol. 360, 11–29. https://doi.org/10.1016/j.ydbio.2011.09.005 (2011).
Harland, R. M. In situ hybridization: an improved whole-mount method for Xenopus embryos. Methods Cell Biol. 36, 685–695. https://doi.org/10.1016/s0091-679x(08)60307-6 (1991).
Takebayashi-Suzuki, K., Arita, N., Murasaki, E. & Suzuki, A. The Xenopus POU class V transcription factor XOct-25 inhibits ectodermal competence to respond to bone morphogenetic protein-mediated embryonic induction. Mech. Dev. 124, 840–855. https://doi.org/10.1016/j.mod.2007.09.005 (2007).
Acknowledgements
We would like to thank the National Xenopus Resource (RRID: SCR_013731) at the Marine Biological Laboratory (MBL) for technical support of CRISPR/Cas9-mediated KO experiment and the National Bio-Resource Project (NBRP) of the Japan Agency for Medical Research and Development (AMED) for providing X. tropicalis. We are grateful to Dr. Shinpei Imori and Dr. Ryoya Oda at Hiroshima University for their support on statistical analysis, our lab members, Dr. Yutaka Kikuchi, his lab members, and other colleagues for discussion regarding this work. This work was supported by Grant-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology (MEXT), Japan: Japan Society for the Promotion of Science (JSPS) KAKENHI, 19K07247, 20K07225 and 23K06304 to A.S. and K.T.-S. This work was also supported by Hiroshima University Natural Science Center for Basic Research and Development, by Japan Science and Technology Agency (JST) SPRING, JPMJSP2132 to M.N. and R.K., and by National Institutes of Health (NIH) P40 OD010997 and R24 OD 030008, United States, to M.E.H. We thank the Hoansha Foundation for their generous financial support.
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M.N. and A.S. designed and directed the project; M.N., T.K., H.Y., K.T.-S., R.K., E.T., and Y.M. performed the experiments; H.Y., K.T.-S., M.W., and M.H. were involved in the planning and supervised the work; M.N. and A.S. took the lead in writing the manuscript. All authors provided critical feedback and helped to shape the research, analysis, and manuscript.
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Nakamura, M., Kyoda, T., Yoshida, H. et al. Injury-induced cooperation of InhibinβA and JunB is essential for cell proliferation in Xenopus tadpole tail regeneration. Sci Rep 14, 3679 (2024). https://doi.org/10.1038/s41598-024-54280-w
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DOI: https://doi.org/10.1038/s41598-024-54280-w
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