Whole body regeneration deploys a rewired embryonic gene regulatory network logic

Domain: neuroscience
Published: 2025-12-11
Editorial outcome: major_revision

Abstract
A long-held hypothesis in regeneration proposes that developmental processes are re-deployed during regeneration. To investigate this, we compared embryonic and regeneration gene regulatory networks (GRN) in the sea anemone Nematostella vectensis using transcriptomic time series spanning these two developmental trajectories. Here, we show that regeneration reuses cohorts of the embryonic genes along with a small set of genes whose expression dynamics are specific to regeneration. We identified co-expression modules that are either conserved between embryogenesis and regeneration or specific to regeneration, with the latter linked to cellular mechanisms such as apoptosis, tissue remodeling, and wound healing. Functional assays revealed that apoptosis and cWnt signaling pathways are partially MEK/ERK dependent, have largely distinct downstream targets but converge to coordinate regenerative responses. Collectively, these results indicate that regeneration in N. vectensis represents a partial redeployment and extensive rewiring of the embryonic GRN, reactivating developmental modules through a regeneration-specific network logic.

Introduction
Regeneration of cells, tissues, appendages, or even entire body parts is a widespread yet still a rather poorly understood phenomenon in the animal kingdom1. A long-standing question in the field of regeneration is whether and to what extent embryonic gene programs, initially used to build an organism, are reused during regeneration2,3. Several transcriptomic studies of regeneration in axolotl, anole, zebrafish, and sea anemones have highlighted the importance of the re-deployment of developmental pathways4–10. Many studies have directly compared embryonic and regenerative gene expression, focusing on single or groups of genes, revealing (i) genes that are specific to embryonic development11, (ii) genes that are specifically expressed or required during regeneration12,13, and (iii) embryonic genes that are reused during regeneration to some extent14–18.

One study has addressed the question by comparing the transcriptional dynamics between post-larval development and regeneration from dissociated cells in sponges, further highlighting partial similarities between these two developmental trajectories19. Another study has investigated the relationship between larval skeleton development and brittle star arm regeneration with a special emphasis on FGF signaling, revealing that a skeleton developmental gene regulatory module is re-deployed during regeneration20. Investigating the morphological, cellular, and transcriptomic relationship between leg development and regeneration in a marine arthropod (P. hawaiiensis), the authors described (i) that regeneration produces an exact replica of the leg21 and (ii) that both processes involve similar sets of genes, although their temporal relationship seems to be different22.

Despite these studies, the extent to which gene regulatory network programs are reused globally during regeneration, as well as the functional comparison between these two developmental trajectories, remains unknown. The sea anemone Nematostella vectensis (Cnidaria, Anthozoa) offers a unique lens into this question as it affords large-scale functional comparisons between embryonic development and whole-body regeneration within the same organism.

Nematostella vectensis (N. vectenis) is a research model in which embryonic development and whole-body regeneration can be studied (Fig. 1A), and thus, ideally suited for this line of inquiry23. N. vectensis has long been used as a model system to study embryonic development, evolution of body patterning, and gene regulatory networks24–30. More recently, N. vectensis has emerged as a powerful whole-body regeneration model as it is capable of re-growing missing body parts in less than a week9,31–39, reviewed in ref. 40. Local injury-induced regeneration in N. vectensis causes a systemic response throughout the entire body that is crucial for maintaining homeostasis during regeneration41. This process follows a early regenerative program (8–20 hpa) and ending with a late regenerative program which approaches the uncut condition (24–144 hpa). Regeneration proceeds through a wound-healing phase (0–4 hpa) followed by the dynamic but highly stereotypical morphological and cellular program (i.e., regeneration stages), involving tissue remodeling and the de novo formation of body structures35,36. More precisely, regeneration requires a Tissue Contact (TC) between the mesenteries (MES) and the body wall epithelia (BWE) at the amputation site37, cell proliferation34,36, as well as two populations of fast and slow cycling, potential stem cells37. It has been suggested that MEK/ERK signaling is required for oral regeneration38, and recently, a study has shown that ROS (Reactive Oxygen Species) signaling is crucial for post-TC regeneration steps by controlling cell proliferation in a tissue-specific manner42. While many developmental signaling pathways are deployed during regeneration9,38,39,41,42, their precise roles as well as their regulatory logic remain unknown.

Here, we take advantage of the biological and experimental properties of N. vectensis to address the historical question to what extent regeneration recapitulates embryonic development and to decipher molecular signatures unique to regeneration. In the present study, we performed a global transcriptomic comparison of embryogenesis and regeneration using deeply sampled transcriptomic datasets. This approach was combined with a pathway-specific functional analysis as well as a comparison of the embryonic and regeneration GRNs within the same species. Overall, this study (i) revealed that whole-body regeneration is transcriptionally modest in comparison to embryonic development; (ii) identified genes, cellular processes and network modules specific to the process of regeneration; (iii) demonstrated the plasticity of the network architecture underlying two developmental trajectories; and finally, (iv) showed that whole-body regeneration deploys a rewired embryonic GRN logic to reform lost body parts.

Results
Regeneration is a partial re-use of embryonic development
To compare embryogenesis and regeneration (Fig. 1A) on a global transcriptome-wide scale we analyzed four RNAseq datasets; one spanning 16 timepoints of regeneration39 and three spanning a total of 29 embryonic timepoints39,43,44 that were batch corrected39. To assess the transcriptomic states underlying embryogenesis we performed principal component analysis (PCA) on batch corrected embryonic data (Fig. 1B). We found that most gene expression changes occur during the first day of embryonic development from cleavage to blastula stage (Fig. 1B, 7 h post fertilization (hpf) – 24 hpf, PC1 proportion of variance 68%; PC2 proportion of variance 20%) indicating large transcriptomic differences in early embryogenesis. From 96 hpf onwards the samples exhibited modest changes in transcriptional variation suggesting that most transcriptional dynamics driving embryogenesis is complete by this stage (96–240 hpf).

When we examined the regenerative program using PCA (Fig. 1C), we observe three distinct transcriptional programs based on the variations in gene expression: (i) a wound-healing phase (0–4 h post amputation (hpa)) that is followed by (ii) the activation of the early regenerative program (8–20 hpa) in which the samples are distributed along the second principal component (PC2 proportion of variance = 24%, Fig. 1C). These two phases can be correlated to morphological and cellular processes observed in earlier studies, i.e., the completion of wound-healing at 6 hpa36 as well as the onset of mitotic activity 12–18 hpa34,36. (iii) From 24 hpa onwards, most of the variation in gene expression is explained by the first principal component during the late regenerative phase (PC1 proportion of variance = 48%, 24–144 hpa, Fig. 1C). This phase corresponds to the burst of cell proliferation as well as the reformation of oral structures such as the pharynx, the tentacles and the mouth36 (iv) Towards the end of regeneration (144 hpa), we observe a transcriptomic profile that resembles the uncut samples indicating a return to steady state. These profiles correlate with the major events of oral regeneration in N. vectensis and indicate that our sampling strategy effectively covers the major transcriptional hallmarks of regeneration36.

We next directly compared the transcriptomic variation of regeneration and embryogenesis using PCA and found that the transcriptional changes during regeneration were relatively modest compared to those observed during embryogenesis with most of the variation in the first two principal components being driven by the embryonic data (PC1 proportion of variance = 67%, PC2 proportion of variance = 16%, Fig. 1D). This indicates that the transcriptional dynamics of embryogenesis are more profound than those of regeneration. This finding was buttressed by comparing the number of ‘dynamically expressed genes’, those which are significantly differentially expressed log₂FC > 1 or < −1 and false discovery rate (FDR) < 0.05 at any timepoint compared to t0 defined as 0 hpa for regeneration and 7 hpf (the onset of zygotic transcription) for embryogenesis. Embryogenesis exhibited more than ten times the number of dynamically expressed genes compared to regeneration (15,610 and 1254 genes, respectively, Fig. 1E). These results further show that regeneration, when compared to embryogenesis, employs far fewer dynamic genes to accomplish a similar task: (re)constructing a functional animal. Of the dynamic genes observed during regeneration, the vast majority (1131 out of 1254) are also dynamically expressed during embryogenesis, suggesting that regeneration is largely a partial re-use of the embryonic gene complement (Fig. 1E).

Identification of genes with regeneration-specific expression dynamics
Among the genes dynamically expressed during regeneration, a small fraction, 123 genes, exhibit significant differential expression (fC > 2-fold) only during regeneration which we term “regeneration specific” (Supplementary Data 1). There is no obvious bias into a specific temporal expression pattern, however, they are enriched in the regeneration clusters39 R2, R3, R6, and R9 (Fig. 1F). Furthermore, 45 of these genes are detectable exclusively during the regeneration process indicating they are transcriptionally silent during embryogenesis. The remaining 78 genes are expressed during embryonic development but are not considered dynamic from our differential expression analysis above (Fig. 1F). Interestingly, several of the 123 “regeneration specific” genes, for example wntless (NvERTx.4.10017) and agrin-like (NvERTx.4.43369), have previously been reported to be important regulators of regeneration in bilaterians45,46. Furthermore, among the “regeneration specific” genes, 47 have no known homology in the Uniprot database (BLASTp, e value cutoff < 0.05, see “Methods” for annotation details), suggesting possible taxonomically restricted genes. These results indicate not only a possible evolutionary conservation of gene use in regeneration but also identify additional genes that may play important roles specifically during whole-body regeneration. A gene ontology (GO) term enrichment analysis on these 123 “regeneration specific” genes, revealed a suite of biological process GO terms relating to modulation of signaling pathways (e.g., wntless), metabolic processes, and apoptotic cell death, indicating an essential role for these processes in regeneration (Fig. 1G).

Embryonic gene modules are partially redeployed during regeneration
While keeping in mind the difference between the two processes, i.e., development of an entire organism vs regeneration of oral structures, it is nonetheless interesting to note that regeneration dynamically expresses less than one-tenth the number of genes compared to embryonic development. We were thus interested in how these genes were deployed and arranged into co-expression networks. We sought to determine if embryonic gene network modules themselves are reused in a reduced capacity or if regeneration deploys novel gene module arrangements. For this, we used previously determined fuzzy c-means gene expression clusters39, i.e., eight embryonic clusters (Figs. S1 and S2A) and nine regeneration clusters (Figs. S1 and S2B).

To explore these expression clusters (also named modules), we performed GO-term enrichment for each cluster (Supplementary Data 2 and 3) and examined the clusters at the gene level. We found that modules that were activated early in either of the two processes (e.g., embryogenesis cluster 4 (E4), regeneration cluster 6 (R6), Fig. S2A, B) contained several canonical developmental genes: wntA (NvERTx.4.132141), lmx (NvERTx.4.163477) and foxA (NvERTx.4.73096) in embryonic cluster 4 (Fig. S2Ai) and tcf (NvERTx.4.100051), spr (NvERTx.4.64399), and runx (NvERTx.4.92298) in R6 (Fig. S2Bi). The early embryonic transcriptional activation of wntA, lmx and foxA in the endomesoderm/central ring has been previously reported26,47–49 and confirmed by in situ hybridization at 24 hpf (Fig. S2Aii). The spatio-temporal expression patterns of tcf, spr and runx during regeneration (uncut, 0–60 hpa, Fig. S2Bii) confirm the dynamic expression patterns of these genes as early as 2 hpa at the amputation site. We conclude from this analysis that classical developmental genes are involved in the early phases of both embryogenesis and regeneration.

We next analyzed whether the same groups of genes were co-regulated during embryogenesis and regeneration, by testing if gene expression observed during both processes was arranged in similar co-expression modules (Fig. 2). We compared regeneration and embryonic clusters on a gene-cluster membership basis to identify significant overlaps. Regeneration clusters with high overlap of a specific embryonic cluster indicate a shared or reused network logic since the same suite of genes is deployed as a bloc in both processes. Regeneration clusters with low overlap to any single embryonic cluster, on the other hand, are likely to be de novo genetic arrangements specific to regeneration.

We found that most of the regeneration clusters exhibited significant overlap with one or more embryonic clusters (Fig. 2A, B). These “conserved modules” also exhibited high preservation permutation co-clustering z-scores (Fig. 2B; >2 indicating conservation; >10 indicating high conservation; permutations = 1000)50. Importantly, we identified two clusters, R1 and R6, which exhibited relatively low overlap with anyone embryonic cluster, indicating that these are likely “regeneration specific” arrangements. When we examined the GO-term enrichment of each cluster, we found that in general, highly conserved clusters (e.g., R5) were enriched in GO-terms corresponding to homeostatic cell processes (Fig. 2C), while lowly conserved regeneration specific clusters (e.g., R6) were enriched in GO-terms describing developmental signaling pathways (Fig. 2F). These results suggest that the gene module arrangements pertaining to core biological functions such as cell proliferation are common to embryogenesis and regeneration while the gene modules containing developmental patterning genes are unique to each process.

Two clusters that exemplify these findings are R5 and R6. Cluster R5, a conserved cluster (zStatistic 6.94) showed strong enrichment of cell-proliferation related GO-terms (Fig. 2C). When we examined exemplar genes (with intra-module membership scores > 0.95) ercc6-like (NvERTx.4.116232), rad54B (NvERTx.4.92456), mcm10 (NvERTx.4.112709), cyclinB3 (NvERTx.4.144958), we observed co-expression patterns during embryonic development (cluster E-1, Fig. 2D) and regeneration (cluster R5, Fig. 2E) further demonstrating module conservation. The expression pattern of these genes during regeneration correlate well to the timing of proliferation activation during N. vectensis regeneration with an activation at 24 hpa, a peak at 48 hpa, and a taper off thereafter34,36 (Fig. 2E).

In contrast, cluster R6 is specific to regeneration. This module exhibits strong enrichment of GO-terms relating to apoptosis and developmental signaling pathways (Fig. 2F). When we examined four exemplar genes tcf (NvERTx.4.100051), bax (NvERTx.4.118493), runx (NvERTx.4.92298), bcl2 (NvERTx.4.81296), we observed divergent profiles during embryogenesis (Fig. 2G) but co-expression during regeneration (Fig. 2H) indicating that this grouping of genes is indeed “regeneration specific”. These results suggest that modules containing genes responsible for basic cellular functions are largely reused and co-expressed between embryogenesis and regeneration, while those including genes that are important for the activation of developmental processes have regeneration-specific arrangements.

Identification of synexpression groups at the onset of morphogenetic movements
We next decided to characterize the potential synexpression groups comprising the GRN underlying whole-body regeneration in N. vectensis at 20 hpa. This timepoint was chosen as it corresponds to the onset of morphogenetic movements, invagination/tissue reorganization36, as well as the likely onset of the regenerative program (Fig. 1C). Thus, this timepoint will allow a comparison with the previously defined embryonic GRN at the onset of the morphogenetic movements of gastrulation26,51.

To identify a set of genes relevant for our study and potentially involved in coordinating the regenerative response we performed a differential gene expression analysis between 0 and 20 hpa and identified 2263 transcripts of which ~1400 were differentially expressed with an at least 2-fold variation (Fig. 3A; Supplementary Data 4). We cross-referenced this list of transcripts with the “regeneration-specific” genes identified above (Supplementary Data 1) as well as with genes that are part of the GRN underlying embryonic development, i.e., 15 embryonic cWnt targets26 (Supplementary Data 5) and 25 embryonic MEK/ERK targets51 (Supplementary Data 6) downstream targets. These embryonic downstream targets are largely pathway-specific as only 4 genes (nfix-like, hd050, elk-like 1, and gsc) are controlled by MEK/ERK and cWnt at the onset of gastrulation (Supplementary Data 5 and 6). Doing so, we identified 10 embryonic GRN (6 MEK/ERK + 4 cWnt downstream targets) and 44 “regeneration-specific” genes that were downregulated at 20 hpa (Fig. S3; Supplementary Data 7, 8, and 9). Furthermore, we identified 30 genes that are part of the embryonic GRN (19 MEK/ERK51,52 and 11 cWnt26 downstream targets) as well as 40 “regeneration-specific” genes that were upregulated at the onset of regeneration (Fig. 3B; Supplementary Data 7, 8, and 9).

Performing an in situ hybridization screen for these genes we obtained 26 patterns that we regrouped based on their spatial expression profiles at 20 hpa (Fig. 3C). Genes were either expressed in the amputation site ectodermis (AE: wnt2, foxB, wntA; Fig. 3Ca–c), amputation site gastrodermis (AG: wnt7b, nkd1-like, wnt4, fz10; Fig. 3Cd–g), amputation site gastrodermis + amputation site mesenterial Tips (AG/AMT: runt, moxD, axin-like, twist, wls; Fig. 3Ch–l), amputation site gastrodermis + mesenteries (AG/M: sprouty, musk-like, porcupine-like, smad4-like; Fig. 3Cm–p), mesenteries (M, hd50, mae-like, bicaudal-like, k50-5, pdvegfr-like, fox1, foxD1, mox1, phtf1-like; Fig. 3Cq–y) and the physa (P, fgfA2; Fig. 3Cz). Taken together with their temporal expression profiles reflected by their expression clusters (Fig. 3B; Supplementary Data 7, 8, and 9), most of these genes form synexpression groups53, potentially involved in the same biological process and underlying gene regulatory networks within their respective expression domains.

Injury-induced regeneration activates apoptosis, pERK, and cWnt
Having observed a strong enrichment for apoptosis related GO-terms in the list of 124 regeneration-specific genes and in the regeneration specific module R6, we investigated the dynamics of apoptosis during the regenerative process. Several genes relating to apoptosis, including the regeneration-specific genes bax (NvERTx.4.118493), caspase-3 (NvERTx.4.152711), bcl2 (NvERTx.4.81296), and an additional bcl2 (which we term bcl2B; NvERTx.4.84383), belong to the regeneration specific cluster R6 and are activated shortly after amputation (Fig. 4A; Supplementary Fig. S4A). Whole mount in situ hybridization for bax 24 hpa confirms the increased dynamics and indicates that like other genes from the expression cluster R6 it is activated at the amputation site as early as 2 hpa and progressively increases its expression in the mesenteries in later stages (Fig. 4C; Supplementary Fig. S4B).

We next performed a time series of TUNEL staining to examine the dynamics of apoptosis during embryogenesis and regeneration. While only few apoptotic cells can be observed during embryonic development54, during regeneration we observed a burst of apoptotic activity after amputation at the cut site as early as 1.5 hpa which perdured through 12 hpa (i.e., 1st apoptotic wave; Fig. 4D; Supplementary Figs. S4 and S5). At 24 hpa apoptotic activity is now randomly detected along the body wall epithelia of the entire polyp, potentially involved in the systemic response (i.e., 2nd apoptotic wave; Fig. 4D; Supplementary Fig. S4). This dynamic TUNEL profile is reminiscent of the bax expression pattern.

In addition to apoptosis, we assessed the activation of MEK/ERK and cWnt signaling pathways during the onset of regeneration. This choice has been made to compare the regeneration GRN with the GRN underlying embryonic development that is driven by MEK/ERK/Erg and cWnt/Tcf signaling26,51,52. erg (NvERTx.4.84016) and tcf (NvERTx.4.100051) are the respective MEK/ERK and cWnt effectors involved in N. vectensis embryonic germ layer formation26,51,52. Analyzing their expression pattern during regeneration, revealed that both genes are up-regulated after wound-healing (Fig. 4E, I) and localized at the TC between the oral part of the mesenteries and the body wall epithelia at the amputation site (Fig. 4G, K; Supplementary Figs. S2 and S6).

Using a monoclonal antibody directed against the phosphorylated/activated form of ERK (pERK) during the time course of regeneration (Fig. 4H; Supplementary Fig. S6), we detected localized pERK staining as early as 1 hpa in the body wall epithelia at the amputation site as well as in the oral tips of the mesenteries (Fig. 4H). After wound-healing (6 hpa, Fig. S6), this staining remains localized as regeneration progresses until 48 hpa (Fig. 4H). A localized signal is detected in the reforming pharynx (72, 96 hpa), after which the signal begins to be re-detected ubiquitously throughout the body (120, 144 hpa), reminiscent of the staining detected in uncut controls (Fig. 4H; Supplementary Fig. S6).

To determine localization of cWnt activation, we developed a transgenic knock-in line in which we fused the fluorescent reporter mCherry in frame with the C-terminal end of the endogenous copy of ß-catenin (Fig. S7). Positive F1, Nvß-catenin::mCherry, were outcrossed and offspring used to assess early cleavage stages enabling us to observe nuclearization of ß-catenin (Movie S1) confirming previous observations25–27,30. We then assessed ß-catenin nuclearization in Nvß-catenin::mCherry polyps during the process of regeneration using an anti-mCherry antibody (Fig. 4L). While no nuclear staining was observed at 1h30 hpa, nuclear staining became detectable at 12 hpa and persisted at least up to 24 hpa at the amputation site. Intriguingly, we did not detect any nuclear staining at 48 hpa and 60 hpa.

Taken together these spatio-temporal expression and activation patterns suggest that MEK/ERK signaling, and apoptosis are activated shortly after amputation and during the process of regeneration, while cWnt signaling is activated only during regeneration after wound-healing is completed.

Regeneration in N. vectensis requires apoptosis, MEK/ERK, and cWnt
To test whether apoptosis, MEK/ERK and/or cWnt are required for regeneration, we treated amputated juveniles during the time course of this process with pharmacological inhibitors reported and confirmed to be efficient in N. vectensis; the pan-caspase inhibitor Z-VAD55 to block apoptosis, the MEK inhibitor U012651,52,56 and iCRT14 to block cWnt signaling57,58 (Figs. S8 and S9).

Previous studies have shown that perturbation of MEK/ERK and cWnt block embryonic development and patterning in N. vectensis26,27,51,52,56. We therefore tested whether apoptosis is a regeneration-specific process or if it was also required for embryonic development. Interestingly, N. vectensis embryos treated continuously with Z-VAD after fertilization developed normally into functional polyps, showing no developmental defect and metamorphosed on time as in controls (Fig. 5A). While Z-VAD treated animals were slightly smaller than controls (Fig. S10), the absence of visible phenotypes during embryonic development and metamorphosis support the idea that apoptosis in N. vectensis is a regeneration-specific process when compared to embryonic development.

We then treated regenerating polyps continuously immediately after amputation with either Z-VAD, U0126 or iCRT14 up to 144 hpa. Although all treatments impaired the regeneration program, there were visible differences (Fig. 5B). Z-VAD treatments prevented the emergence of TUNEL positive (TUNEL+) cells as soon as 1.5 hpa (Fig. S8A) and regeneration was blocked in an early regenerative stage (i.e., modified stage 1), lacking the physical interaction between the fused oral tips of the MES and the BWE at the wound site (Fig. 5Bb). U0126 treatments effectively blocked pERK activation as early as 1 hpa (Fig. S8B) and animals were blocked in an even earlier regeneration stage (stage 0.5), i.e., neither the fusion of the MES tips nor the TC between the MES and the BWE of the wound site were observed (Fig. 5Bc). iCRT14 treated polyps did not display ß-catenin nuclearization at 24 hpa (Fig. S8C) and in line with a later activation of the cWnt pathway described above, treated polyps regenerated up to stage 2 but never formed a pharynx or tentacles (Fig. 5Bd).

Inhibition of MEK/ERK signaling (using U0126) or apoptosis (using Z-VAD) from 0 to 144 hpa both block the regeneration process early on. To gain further insight into the roles of these two pathways we performed waves of inhibitory treatments during various time windows and assessed the resulting phenotypes (Fig. S11). When U0126 treatments were started at 24 or 48 hpa, and lasted up to 144 hpa, regeneration was blocked at stage 2 in most cases (100% n = 75/75 and 75% n = 50/75, respectively) (Fig. S11A). When U0126 was added at 72 hpa or later time points up to 144 hpa, regeneration was completed successfully or slightly delayed in comparison to DMSO treated controls. These results indicate that MEK/ERK signaling is crucial as early as stage 0.5 for initiating a regenerative response.

To dissect the temporal contribution of apoptosis during regeneration, we selectively inhibited each of the three apoptotic waves using time-specific Z-VAD treatments and evaluated regeneration outcomes at 144 hpa (Fig. S11B) following the staging system from ref. 36. Independent inhibition of the first apoptotic wave (0–6 hpa), the second wave (12–48 hpa), or the third wave (48–144 hpa) resulted in a substantial proportion of polyps successfully completing regeneration, i.e., 73% (n = 78), 53% (n = 70), and 53% (n = 68), respectively. While the majority of polyps reached stage 4, some showed defects in tentacle formation—especially when apoptosis was blocked during the second (32%) or third (72%) waves. These findings suggest that while apoptosis during each wave contributes to overall regenerative success, apoptosis during the later stages may play a more critical role in the proper morphogenesis of specific structures, such as tentacles.

MEK/ERK signaling is required for injury-induced apoptosis and cWnt
To gain further insight into the different regeneration phenotypes caused by the treatments, we assessed their effects on earlier hallmarks, such as wound-healing (Fig. 5C). Wound-healing is terminated in about 90% (n = 48) of control polyps by 6 hpa36 (Fig. 5C). While none of the treatments blocked wound-healing, it is nonetheless noticeable that this process is significantly delayed in U0126 treatments (wound is healed at 8 hpa in 90%, n = 36), but significantly accelerated in Z-VAD and iCRT14 treatments, i.e., wound is healed at 4 hpa in 100% (n = 27) and 90% (n = 28), respectively (Fig. 5C). To determine whether this delayed wound-healing is responsible for the arrest in the regeneration process, we compared the effects of U0126 treatments starting at 0 hpa or 8 hpa (i.e., after wound-healing is completed36) on their capacity to reach at least stage 1 at 24 hpa. While most control animals (12 out of 15 cases) reached stage 1 at 24 hpa, U0126-treated animals were consistently blocked at stage 0.5 (start 0 hpa: 14 out 16 cases; start 8 hpa: 12 out of 13 cases) (Fig. S12), indicating that regeneration is blocked independently of the delayed wound-healing process.

TC between the MES and the body wall epithelia BWE at the amputation site is a hallmark of regeneration in N. vectensis and observable at 24 hpa and 48 hpa36. In a similar timing, the initiation (24 hpa) and burst (48 hpa) of amputation-induced cell proliferation occurs at the amputation site34,36. We thus performed inhibition treatments between 0–24 hpa, 0–48 hpa or 24–48 hpa, and assessed their impact on the initiation or maintenance of the TC (Fig. 5D, Supplementary Data 10) and cell proliferation (Fig. 5E). In animals treated with Z-VAD we observed that the initiation of the TC occurred in a similar manner than in controls at 24 hpa (Fig. 5Dd). However, its maintenance was impaired at 48 hpa (Fig. 5De, f). More drastically, inhibition of MEK/ERK prevented not only the initiation of TC at 24 hpa (Fig. 5Dg) but also its maintenance at 48 hpa (Fig. 5Dh, i). In contrast, when cWnt signaling was inhibited TC was simply delayed at 24 hpa and maintained at 48 hpa (Fig. 5Dj–l). Further quantification of the number of cells in S-Phase (EdU+ cells) in these same conditions, revealed a significant down-regulation of cell proliferation at the end of all treatment windows (Fig. 5D, E and Supplementary Data 11).

Taken together, these data indicate that injury-induced initiation and maintenance of cell proliferation depends on a constructive function of apoptosis as well as MEK/ERK and cWnt signaling, while the impact of these pathways on the initiation and maintenance of the TC is pathway dependent. Based on this global phenotype assessment, MEK/ERK signaling emerged as to have the earliest impact with the broadest range of phenotypes linked to the initiation of the regeneration program in N. vectensis (Fig. 5F). These observations further indicate that the premature (stage 0.5) arrest of regeneration upon inhibition of MEK/ERK occurs through the inhibition of morphogenetic movements during the proliferation-independent phase of regeneration34,36.

We next sought to understand the potential relationship between these three signaling pathways at 20 hpa (Fig. 6). While Z-VAD treatments had no visible effects on TUNEL staining or the activation of pERK, nuclearization of ß-catenin appeared slightly increased compared to control conditions (Fig. 6A). Following U0126 treatments, all three pathways seemed impaired which was reflected by the absence of TUNEL+ cells, pERK staining and nuclearization of ß-catenin (Fig. 6A). Finally, iCRT14 prevented nuclearization of ß-catenin, did not affect pERK staining but increased the amount of TUNEL+ cells (Fig. 6A). Taken together, these results suggest that MEK/ERK signaling is upstream of apoptosis and cWnt signaling and that apoptosis and cWnt signaling are required for mutual repression during the onset of regeneration (Fig. 6B).

MEK/ERK, cWnt and apoptosis have largely distinct downstream targets
To gain further insight into the molecular mechanisms activated by MEK/ERK, cWnt or apoptosis at the onset of regeneration, we performed bulk RNAseq for each of these treatments at 20 hpa (Fig. 7A and Supplementary Data 12) in adults that display the same phenotypes as juveniles (Fig. S13 and Supplementary Data 13). We assessed significantly differentially expressed genes (log₂FC > 1 or > −1 and an adjusted p < 0.05) in any treatment compared to control/DMSO-treated regenerating animals. We identified 342 (193 up/149 down), 578 (304 up/274 down) and 472 (220 up/252 down) genes that were differentially expressed in Z-VAD, U0126 and iCRT14-treated animals, respectively (Fig. 7B; Supplementary Data 14, 15, and 16).

Strikingly, GO-term enrichment analysis of these differentially expressed genes (Fig. 7C and Supplementary Data 17) largely confirmed the observations made following the experiments to address the functional relationships between these pathways. In line with the observation that MEK/ERK signaling controls cWnt signaling and apoptosis, GO-terms enriched in the dataset of genes down-regulated by U0126 at 20 hpa contained Wnt signaling and apoptosis. Furthermore, and in line with the observation that cWnt and apoptosis are required for mutual repression during the onset of regeneration, GO-terms enriched in the dataset of genes up-regulated by Z-VAD or iCRT14, contain Wnt signaling and neuron death, respectively (Fig. 7C and Supplementary Data 17). Assessing the temporal expression patterns of the genes down- or up-regulated following the various treatments (Fig. S14) further support the central role of MEK/ERK signaling during the onset of regeneration. In fact, while most of the apoptosis and cWnt downstream targets are mainly up-regulated either shortly after amputation or late during the regeneration process, a large part of MEK/ERK downstream targets are also activated at the onset of regeneration, i.e., 12–24 hpa (Fig. S14).

We then compared the lists of differentially expressed genes and observed that MEK/ERK, cWnt and apoptosis during regeneration in N. vectensis appear to have largely distinct downstream targets (Fig. 7D; Supplementary Data 18, and 19). In fact, only a small number of down-regulated genes were shared between the different treatment conditions. Specifically, five genes were commonly downregulated in all three treatments. When comparing pairs of treatments, eight genes were shared between U0126 and Z-VAD, 20 genes between U0126 and iCRT14, and 28 genes between Z-VAD and iCRT14 (Fig. 7D and Supplementary Data 18). Similarly, only a limited number of up-regulated genes were shared across treatments. Specifically, 10 genes were commonly upregulated in all three treatments. Pairwise comparisons revealed 14 shared genes between U0126 and Z-VAD, 39 genes between U0126 and iCRT14, and 45 genes between Z-VAD and iCRT14 (Fig. 7D and Supplementary Data 19).

The absence of strong overlap between the downstream targets of all three pathways might appear surprising because UO126 treatments affect both apoptosis and cWnt activation (Fig. 6Ac, k). The absence of overlap between U0126 and iCRT14 can be explained by the delayed TC in iCRT14-treated animals (Fig. 5Dj), the presence of compensatory regulatory mechanisms that drive cWnt target genes during regeneration in N. vectensis independently of MEK/ERK signaling, or spatially restricted actions of MEK/ERK affecting injury-induced cWNT. The absence of overlap between U0126 and Z-VAD can be explained by the caspase-independent activation of apoptosis at 20 hpa (Fig. 6Ab). Taken together, these data suggest that although MEK/ERK is required for apoptosis and cWnt signaling at 20 hpa, these three pathways seem to act at different moments in a complementary manner during the regeneration process in N. vectensis.

MEK/ERK regulates “embryonic” and “regeneration-specific” genes
We next sought to understand to what extent either of these three pathways are controlling injury-induced expression of the 15 embryonic cWnt (emb_cWnt) and 25 MEK/ERK (emb_ERK) downstream targets as well as the 84 regeneration-specific (regen_spe) genes up- or down-regulated at 20 hpa (Fig. 3). Systematically comparing these pools of genes (emb_cWnt, emb_ERK, regen_spe) with the up- or down-regulated genes following inhibition of MEK/ERK (Fig. 7; Supplementary Data 20 and 21), apoptosis (Fig. 7; Supplementary Data 22 and 23), or cWnt (Fig. 7; Supplementary Data 24 and 25) during regeneration, we observed that MEK/ERK signaling during regeneration has the most widely extended control on gene expression for all three pools (Fig. 8 and Supplementary Fig. S15). This is mostly visible in the down-regulated genes (Fig. 8A). In fact, no genes of the “embryonic” pools are affected by neither Z-VAD nor iCRT14, and only three genes from the regeneration specific genes are downregulated by iCRT14. However, six, seven, and eight genes belonging to the embryonic ERK, embryonic cWnt or regeneration-specific gene pools, respectively, are downregulated by U0126 treatments at 20 hpa (Fig. 8A and Table 1).

To further confirm that these above-identified genes are downstream targets of MEK/ERK during regeneration, we treated animals continuously after sub-pharyngeal amputation or starting at 8 hpa (after wound-healing) with U0126 and analyzed the expression levels of the three gene pools by RT-qPCR at 20 hpa (Fig. 8B; Supplementary Fig. S16). 19 out of the 25 identified embryonic MEK/ERK downstream targets are downregulated by U0126 treatments during regeneration (Fig. 8B). Similarly, 11 out of the 15 identified embryonic cWnt downstream targets are also downregulated following U0126 treatment (Fig. 8B). Finally, for the “regeneration-specific” pool of genes we only assessed 26 out of the 40 identified genes of which 19 were downregulated when MEK/ERK signaling was blocked by U0126 (Fig. 8B). This analysis not only largely confirms but also extends the list of the above described MEK/ERK downstream targets identified by RNAseq (Fig. 8A and Table 1). Furthermore, we also assessed the effects of MEK/ERK inhibition on erg and tcf (insets Fig. 8B), two transcription factors that are effectors of MEK/ERK and cWnt, respectively, during embryonic development in N. vectensis26,51. Interestingly, we observed a reduced expression for erg, suggesting a negative feedback loop, and a strong down-regulation of tcf under those conditions.

In situ hybridization of uncut, 20 hpa DMSO treated and U0126 treated juveniles for a subset of these genes (sprouty, runt, mae-like, erg; twist, foxB, axin-like, tcf; carm1, cad, egln1, and bax), confirmed the RT-qPCR data (Fig. 8C). This analysis also revealed that at the onset of regeneration, MEK/ERK signaling controls expression of genes in several domains of the amputation site, including the AE, AG, AMT and M. Combining the obtained data from RNAseq, RT-QPCR (i.e., temporal expression), and in situ hybridization (i.e., spatial expression) as well as functional studies together, these results enabled us to propose a MEK/ERK-dependent GRN blueprint underlying whole-body regeneration in N. vectensis (Fig. 8D). This GRN blueprint indicates the potential director indirect downstream targets of MEK/ERK signaling at the onset of regeneration, that (i) belong either to the group of embryonic cWnt or embryonic MEK/ERK targets (including their respective potential effector genes tcf and erg) and (ii) are interconnected with “regeneration-specific” genes.

Methods
Animal culture, spawning, embryo rearing, and amputation
Adult N. vectensis were cultured at 16°C in the dark in 1/3 strength artificial seawater (⅓ASW) as previously described87. The day before spawning animals were fed with oysters and were then transferred to a light table for 12 h. Embryos were cultured at 18°C in the dark in ⅓ASW until desired timepoint. Regeneration experiments were performed using 6 weeks-old juveniles or more than 2-month old sub-adults raised at 22°C in the dark in ⅓ASW as previously described36. Gender was not taken into account in this study as males and females are morphologically indistinguishable at these stages.

Pharmaceutical drug treatments
Apoptotic cell death was blocked using the pan-caspase inhibitor Z-VAD-FMK (named Z-VAD throughout the manuscript, #ALX-260-020-M001, Enzo Life Sciences Inc, Farmingdale, NY, USA). A stock solution at 10 mM in DMSO was prepared for Z-VAD, kept at −20°C and diluted in ⅓ASW at a final concentration of 10 µM or 50 µM prior to each experiment (Fig. S4). Each Z-VAD treatment was performed in a final volume of 500 µl in a 24 well plate using the adequate controls. Reagents were changed every 24 h to maintain activity for the duration of the experiments. Phosphorylation/activation of ERK was prevented using the MEK inhibitor U0126 (Sigma-Aldrich U120-1MG). A stock of 10 mM in DMSO was prepared and stored at −20°C. Each treatment with U0126 were performed with a final concentration of 10 μM in ⅓ASW. Treatments were carried the same way as Z-VAD treatments. cWnt activation was prevented using iCRT14, a β-catenin-responsive transcription inhibitor (Sigma-Aldrich - SML0203-5MG). A stock of 10 mM in DMSO was prepared and stored at −20°C. Each treatment with iCRT14 was performed with a final concentration of 20 μM in ⅓ASW. Treatments were carried the same way as Z-VAD & U0126 treatments.

Sample collection and RNA extraction
Sample collection and RNA extraction for the embryonic and regeneration time series are described in refs. 39,88. For pharmacological treatment experiments (biological triplicates), adult animals were first induced to spawn, then amputated below the pharynx and incubated for 20 h in one of the following treatments: 0.1% DMSO (control), 10 µM U0126, 50 µM Z-VAD, or 20 µM iCRT14. At 20 hpa, oral tissues from 10 individuals per condition were dissected and placed into RNAlater (ThermoFisher; #AM7021), transferred to TRI-Reagent (Sigma-Aldrich; #T9424) in Precellys tubes (Bertin; #CK14), lysed (2×0.5 s), flash-frozen in liquid nitrogen, and stored at −80°C. RNA was purified using a double TRI-Reagent extraction with Phase Lock Gel tubes (QuantaBio; #10847-802) and resuspended in 20 µL of nuclease-free water. Potential DNA contamination was removed with TURBO DNase (2 U, 37°C, 25 min) and inactivated at room temperature (5 min). RNA concentration and integrity were confirmed with an Agilent Bioanalyzer; all samples had RIN values > 8.

Library preparation and sequencing
CDNA libraries were prepared by the IRCAN Genome facility using the TruSeq Stranded mRNA Library Prep Kit (Illumina #NP-202-1001) or the Illumina NextSeq 500/550 High-Output v2.5 kit (#20024906). Libraries were sequenced on the Illumina NextSeq 500 platform (75 bp single-end reads).

Read processing, mapping, and quantification
Raw reads were demultiplexed using bcl2fastq and quality-filtered using Trimmomatic89, fastp90, and Cutadapt91. Processed single-end reads were aligned to either a combined transcriptome using Bowtie292 or to the genome (JGI/Nemve193) using STAR94. Transcript-level quantification was performed using RSEM95 for transcriptome-aligned data, and HTSeq96 for genome-aligned data. Gene-level counts were aggregated by grouping transcripts with the same top BLASTn hit to Nemve1 filtered gene models. Gene models that did not have >5 counts in at least 25% of the samples, were excluded. Normalization was conducted independently for each dataset using the edgeR package97. Count-per-million (CPM) values were calculated, and batch effects across embryonic datasets were corrected using the ComBat function from the SVA package98, with developmental timepoint as a categorical covariate. For all downstream calculations only embryonic timepoints from 7 hpf (the estimated onset of zygotic transcription) onward are considered so that only gene transcription dynamics are measured and compared.

Differential expression analysis
Differentially expressed genes (DEGs) were identified using DESeq299 for pharmacological treatment samples, with DMSO 0.1% as the reference. Genes with an absolute log₂FC > 1 or > −1 and an adjusted p < 0.05 were considered significant. For time-series datasets (Fischer, Helm, Warner, and regeneration), differential expression was calculated using edgeR by comparing each timepoint to its respective baseline (t₀: 7 hpf for Fischer and Helm; 24 hpf for Warner; and 0 hpa for regeneration). Genes were classified as significantly differentially expressed if they had an absolute log₂FC > 1 or < −1 and false discovery rate (FDR) < 0.05. These differentially expressed gene lists were compared to identify overlapping and regeneration specific genes. Venn diagrams to visualize overlapping DEGs across conditions were generated using the VennDiagram R package100.

Identification of “regeneration-specific” genes and GO-term analysis
Differentially expressed gene lists were compared to identify overlapping and regeneration specific genes. GO term enrichment were calculated using a Fisher’s exact test and the R package topGO on the GO terms identified from comparing the Nemve1 gene models to the UniProt databases Swissprot and Trembl using the BLASTx like program, PLASTx (e value cutoff 5e-5)101. All identified GO terms were used as a background model. The resulting GO term list was reduced and plotted using a modified R script based on REVIGO102.

PCA, Fuzzy c-means clustering and cluster conservation
The ensuing analyses were performed using log2 (cpm+1) transformed gene-level quantification (Nemve1 filtered gene models). The expression profiles for each Nemve1 gene model were clustered using the R package Mfuzz103 on the combined embryonic dataset and the regeneration dataset separately. The cluster number was set to 9 for the regeneration data and 8 for the embryonic datasets, as these numbers produced well-separated clusters with minimal overlap (Fig. S3) and represent the inflection point at which the centroid distance between clusters did not significantly decrease with the addition of new clusters (Fig. S3). Genes that did not have a membership score above 0.75 were considered noise and designated as cluster 0. Cluster overlap was calculated for genes that were detectable in both datasets using the function overlapTable from the R package WGCNA, using the regeneration cluster assignments as the reference set. A zStatistic of cluster preservation was also calculated using the function coClustering.permutationTest from the WGCNA package104 using the regeneration cluster assignments as the reference set and 1000 permutations.

RT-qPCR
RNA Extraction and RT-qPCR were performed following the protocol from ref. 26. One hundred and sixty juveniles were used per replicate, and each experiment was performed in three biological replicates. We used a 7900HT Fast Real-Time PCR System with 384-Well Block Module (Applied Biosystem) with Faststart universal SYBR Green Master (rox) (FSUSGMMRO Roche). The relative fold change for each gene expression was calculated by comparing treated and untreated conditions, where gapdh or actin was used to normalize fold change (FC).

Animal fixation. After relaxing N. vectensis polyp in MgCl 7.14% in ⅓ASW for 10–15 min, animals where fixed as followed: (i) for Whole mount in situ hybridization in cold Glutaraldehyde 0.2% (25% stock Electron Microscopy Sciences; #16216) + paraformaldehyde 4% (32% stock Electron Microscopy Sciences; #15714) in ⅓ASW for 2 to 5 min on ice, then with paraformaldehyde 4% in ⅓ASW for 1 h on ice. Fixed animals were washed five times in PBTw 0.1% (PBS 1x + Tween 0.1%) in nuclease free water (NFW), then twice with MeOH 100% and stored at −20°C; (ii) for Immunostaining and other stanning in 4% paraformaldehyde (Electron Microscopy Sciences; #15714) in ⅓ASW during 1 h at 22°C or overnight at 4°C. Fixed animals were washed three times in PBT 0.5% (PBS 1x + Triton 0.5%) in NFW and stored at 4°C.

Whole mount in situ hybridization
Whole mount in situ hybridization was performed as previously described105. 0.5–2 kb digox-igenin-labeled (Roche, #11573152910) riboprobes were synthesized using the MegaScript Transcription Kit (Ambion). Hybridization of riboprobes (1 ng/ml diluted in fresh hybridization solution) was carried out at 62°C in 50% formamide hybe buffer and visualization of the labeled probe was performed using NBT/BCIP as substrate for the alkaline phosphatase-conjugated anti-DIG antibody (AP-anti-Dig, Roche, #11093274910). AP/Anti-Dig was used at 1:5000 in blocking solution and incubated at 4°C overnight. In situ hybridization images were taken on a Zeiss Z-1 Axioimager mounted with an Axiocam camera triggered by the Zen software (Carl Zeiss).

Immunostaining
The phosphorylated form of ERK was visualized using a monoclonal antibody directed against its active form, pERK (Cell Signaling Technology; #4377) and the protocol used was described in ref. 38. To be able to visualize the nuclearization of ß-catenin::mCherry (i.e., the activated form) we used the primary anti-mCherry antibody (dilution: 1/200, Abcam; #ab167453), an Alexa Fluor 488 goat anti-rabbit secondary antibody (dilution: 1/200, Invitrogen; #A11008) and followed the protocol described in ref. 106.

Hoechst and Phalloidin staining
For morphologically characterizing the regeneration step, we counterstained the nuclei and actin filament with Hoechst (Invitrogen; #33342) and BODIPY FL PhallAcidin 488 (Molecular Probes; #B607), respectively. Hoechst was diluted to 1/5000, and BODIPY FL PhallAcidin 488 was diluted to 1/200 in PBS 1X.

EdU staining
For cell proliferation analysis, we used the Click-iT EdU kit (Invitrogen; #C10646, #C10337 or #C10339) following the protocol from ref. 34 with the following change: Juveniles were starved for a week to remove homeostasis proliferation and labeled with 5’-ethynyl-2’-deoxyuridine (EdU 0.1 mM in ⅓ASW) for 1 h, then washed 3 times in ⅓ASW prior fixation and EdU revelation.

Apoptotic cell death staining
To detect cell death the “In Situ Cell Death AP kit” (Roche; #11684809910) was used. The manufacturer protocol was modified as described as follows: (1) Fixed animals were permeabilized using 0.01 mg/ml Proteinase K for 20 min at 22°C; (2), Washed twice in PBS 1x; (3), Refixed in 4% paraformaldehyde in PBS 1x for 1 h at 22°C; (4) Washed 5 times in PBS 1x; (5) Incubated with 50 μL of TUNEL reaction mixture for one hour (Roche protocol); (6) Washed 5 times in PBS 1x; (7) Fixed animal were observed for 488 fluorescence. Positive (DNase I treatment after step 4) and negative (without TUNEL-Enzyme) controls were obtained using the manufacturer’s “In Situ Cell Death AP kit” protocol.

Generation of a Nvß-catenin::mCherry KI line
To generate the Nvß-catenin::mCherry KI line, we used a short-homology arm approach described in refs. 107,108 and CRISPR/Cas9 genome editing performed following the protocol described in ref. 109. Briefly, sgRNAs targeting the sequences indicated below were determined using the ZiFiT Targeter website (http://zifit.partners.org/ZiFiT_Cas9) and cloned into the pDR274 vector to create a gRNA expression vector for each target site of interest110. sgRNAs were transcribed using the T7 Megascript kit (Invitrogen; #A57622) and cleaned with the Megaclear kit (Invitrogen; #AM1908). To obtain the donor template, mCherry was amplified using primers that contained short mCherry sequences and short s homology arms (~70 bp). The two sgRNA and repair template were co-injected together with Cas9 into fertilized and dejellied zygotes following the protocol described in ref. 111. Founder F0 polyps expressing the transgene in a mosaic manner were determined visually and raised to sexual maturity and outcrossed to obtain heterozygous F1 (Supplementary Fig. S7B). Offspring of these F1 were used for experimentation.

sgRNA target sequences
sgRNA1: 5’-CCAGCCAAGTGTTCATC-3’
sRNA3: 5’-GGAGGACACTACCAAAAT-3’
Homology arms (lowercase letters indicate mCherry sequences)
FWD_homology_arm: 5’ CATGGCGACCCGAACATGAGTCATCAGC CACCGGGAGGACACTACCAAAATCCAGGGGGTCCTCTGTATGACAC AGAC atggtgagcaagggcgagga 3’ REV_homolgy_arm: 5’ CTTTTGACGTTG CAATCAACACCCAGCTTTGGTGATGAACACTTGGCTGGCGGCCAAAA GGGGGGTGTGGATTAT ttacttgtacagctcgtcca -3’.
Genotyping primers
Fwd1: 5’-CATGGACACGTACCAGATGC-3’
Rev1: 5’-TACAAAATTCGCGAGAACGA-3’

Protein extraction and western blotting
The protein extracts were obtained from 15 adults per replicate and each experiment was performed in triplicate. The animals were placed in 1.5 ml Eppendorf tubes and spun on bench centrifuge to remove all 1/3 ASW before adding 300 μl of lysis buffer (HEPES 50 mM, NaCl 150 mM, NaF 100 mM, EDTA 10 mM, NA4P2O7 10 mM). Then animals were sonicated 5×10 s with incubation on ice between each round. After sonication 200 μl of Lysis buffer complemented with 1% Triton X-100 and a protease inhibitor cocktail (Apoprotine 20 μg/ml, Vanadate 1 mM, AEBSF 250 μg/ml, Leupeptine 5 μg/ml) was added. The samples were then centrifuged for 20 min at 13,500×g at 4°C before collecting the supernatant into new tubes. The BC assay protein quantification kit (Interchim Upima, 40840A) was used to quantify the protein concentration. Samples were subsequently aliquoted by mixing 75 μl of protein extract with 25 μl of 4X Laemmli buffer (Bio-Rad #1610747) and stored at −20°C. Electrophoresis was carried out in 7.5% SDS polyacrylamide gel and samples were denatured at 85°C for 5 min before loading. For each sample, we used 30 μg of protein and followed the protocol described in ref. 18. Briefly, migration was carried in migration buffer (Tris 3 g/L, Glycine 14.2 g/L, SDS 1%) and transfer to the nitrocellulose membrane was performed in transfer buffer (Tris 3 g/L, Glycine 14.4 g/L, 20% Ethanol). Then the nitrocellulose membrane was saturated with Salin buffer (Tris 0.24 g/L, NaCl 1.63 g/L, 5% cBSA, 0.5% Tween-20) and incubated 1 h at room temperature with 1/2500 of primary antibody anti-pERK (Cat. #4377; Cell Signaling Technology) diluted saline buffer. Revelation was carried out by chimioluminescence (EMD Millipore™ Substrats of chimioluminescence HRP Western Luminata™) on a chemioluminescence Imaging – Fusion SL (Vilber).

TOPFLASH assays
Reporter gene assays were performed based on ref. 112. The TOP Flash reporter plasmid (Addgene; plasmid #12456) was amplified in competent E. coli and purified using standard plasmid extraction methods. The injection mix was prepared by diluting the plasmid to 25 ng/µL in nuclease-free water containing 1× Dextran Green 488 as a tracer. The plasmid solution was microinjected into freshly fertilized and de-jellied N. vectensis embryos. Following injection, embryos were washed and incubated overnight at 17°C in one of the following conditions: 0.1% DMSO (control), 10 µM U0126, 20 µM iCRT14, or a combination of 10 µM 1-Azakenpaullone (AZ; Sigma Aldrich A3734) and 20 µM iCRT14. Embryos were collected at 24 hpa for analysis. Luciferase activity was measured using the Luciferase Assay System (Promega; #E1500). Embryos (~100 per replicate) were lysed in 100 µL of lysis reagent, and the lysate was centrifuged to collect the supernatant. For each measurement, 20 µL of lysate was combined with 100 µL of assay buffer, and luminescence was immediately recorded using a Centro XS3 LB 960 luminometer (Berthold Technologies) with MikroWin 2000 software. Three biological replicates were performed per condition, each of which was measured in technical triplicates.

Statistical analysis. Prior to inferential testing, data distributions were assessed for normality using the Shapiro–Wilk test, and for homogeneity of variances using Bartlett’s test. These tests were conducted independently for each dataset and timepoint. For wound-healing assays, since the data were categorical, pairwise Fisher’s exact tests were used to compare each experimental condition against the DMSO-treated control at each timepoint. For EdU incorporation data, following normality and variance assessments, pairwise Wilcoxon–Mann–Whitney tests (non-parametric) were applied to compare each treatment to the DMSO control at each timepoint, as data were not normally distributed. For luciferase assay, following normality and variance assessments, Kruskal–Wallis test (non-parametric) was applied. To identify specific group differences, post-hoc comparisons were conducted using Dunn’s test with a Benjamini–Hochberg (BH) correction for multiple testing.

All statistical analyses were conducted using R (version 4.3.2). A significance threshold of p < 0.05 was applied throughout. Statistical results are presented in Supplementary Data 26. Throughout the manuscript, n = number of biological samples.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript of Johnston et al. compares the transcriptional program driving Nematostella head regeneration to that of its embryonic development. The authors use previously published RNA-Seq data (Warner et al., 2018; Fischer and Smith, 2013; Helm et al., 2013) to show that transcriptional changes during regeneration are relatively small in comparison to the transcriptome dynamics during embryonic development – a finding, which, in my view, is not too surprising. Transcriptome analyses allowed the identification of genes, whose expression was required for the regeneration process. Some of these genes appear to only be involved in the regeneration but not in the embryogenesis program (although some genes on that list are problematic). Johnston et al. suggest, in line with many other studies in different models and using different methods, that Nematostella head regeneration re-deploys parts of the embryogenesis program, but with some regeneration-specific features. Importantly, among the genes upregulated during regeneration, the authors find several key regulators of apoptosis and show that apoptosis is required for successful regeneration but not for the embryonic development. Finally, they address the role of MEK/ERK signaling in the regeneration process. The authors show that wound healing does not depend on MEK/ERK signaling but the onset of regeneration does, and characterize the effect of inhibition of MEK/ERK by QPCR and ISH on a number of genes expressed in different regions of the regenerate.\nI think that this paper has to be significantly improved to reach the Nature Communications standard. In its present state, it is a descriptive transcriptomic study with two side-stories (functional analyses of apoptosis and MEK/ERK signaling during regeneration), which are not interconnected. However, the manuscript has a clear potential to become excellent if the authors dig deeper into their important functional findings. Since activation of apoptosis seems to be a unique prerequisite for regeneration, I think, this should be the focus of the paper. However, Johnston et al. need to find out what triggers apoptosis and what is being triggered by it.\nTo that end, I suggest:\ni) comparing transcriptomes of the ZVAD-treated, U0126-treated and control regenerates at stages 0.5 and 1, identifying the overlap among the regulatory genes significantly affected by the treatments, and characterizing their expression during regeneration;\nii) checking whether apoptosis, MEK/ERK signaling, and expression of b-catenin signaling targets are induced by wounds not causing tissue loss (punctures, incisions), and during foot regeneration;\niii) checking the effect of pharmacological upregulation and downregulation of b-catenin signaling and MEK/ERK signaling on apoptosis at the onset of regeneration;\niv) checking the effect of pharmacological upregulation and downregulation of b-catenin and MEK/ERK on cell proliferation at the onset of regeneration;\nv) checking the effect of ZVAD on pERK (immunohistochemistry) and expression of b-catenin target genes (ISH) in amputated animals;\nvi) attempting to rescue the effect of the ZVAD-mediated inhibition of apoptosis on regeneration by activating b-catenin signaling and/or MEK/ERK signaling.\nAmong all the proposed treatments, only upregulation of MEK/ERK seems not to have been routinely used in Nematostella. However, since FGFRb is expressed in the oral region of Nematostella, recombinant FGF incubation appears to be a plausible way to achieve upregulation of MEK/ERK.\nOnce the authors have experimentally analyzed the links and clearly established the regulatory hierarchy between apoptosis, proliferation, MEK/ERK signaling, and Wnt/b-catenin signaling, and identified and characterized the candidate genes responsible for the start of regeneration, the article can be re-considered for publication at Nature Communications.\nIn addition to the general suggestions above, I list specific things for the authors to address in the revision (in order of appearance in the text, not in order of importance).\n1. Lines 61-62. Johnston et al. write about genes “specifically expressed or required during regeneration” as opposed to “genes specific to embryonic development” and “re-used” genes, and refer to a paper showing the involvement of Sox2 in the regeneration but not in the development of the hair follicle. This is misleading – Sox2 is regeneration-specific only in the hair follicle context, but it is a crucial regulator of the embryonic development.\n2. Lines 93-98. Johnston et al. certainly addressed i) and ii). I am not sure about how the “plasticity of the network architecture” (iii) was demonstrated, and I do not think that the paper shows anything with regard to the “rewiring” (iv). If Fig. 7B is meant, I do not find that this paper convincingly proves experimentally that MEK/ERK is upstream of Wnt/b-catenin signaling during regeneration. If the authors refer to lines 344-362, I am not sure I see evidence for rewiring there: most of the embryonic MEK/ERK targets are still MEK/ERK targets during regeneration.\n3. Lines 117-122. How exactly are the phases defined? If by transcriptomic changes, there probably should be some metric to it. How do they correspond to the morphological phases of Amiel et al., 2015?\n4. Lines 152-154. How exactly do the authors define “regeneration-specific genes”? I did not check all the 124 genes, of course, but I am sure one cannot call Wntless a “regeneration-specific gene”. According to the authors’ own NvERTx database, Wntless is a maternally deposited transcript, and Wntless gene is strongly expressed throughout the development, which is absolutely not surprising, given the importance of Wnt signaling in development and the fact that Wntless is a critical member of the Wnt secretion pathway.\n5. Lines 241-243. The finding of the requirement of apoptosis for regeneration but not for normal development is among the most interesting things reported in the paper. Currently, the results of TUNEL on embryos are “data not shown”. Please show and quantify these data and add them to Fig. 4Bii! Please provide quantification data (Fig. 4Bii) in a form of box plots or individual datum points rather than a table with means, so that it is possible to estimate the variance between polyps. I also suggest increasing N to at least 5 in every case.\n6. Lines 259-260. The authors suggest that apoptosis induces cell proliferation. In order to make the argument that apoptosis is upstream of proliferation more convincing, I suggest directly blocking proliferation (with cyclin-dependent kinase or DNA polymerase inhibitors) and checking whether apoptosis is still detectable.\n7. Line 270. The authors have to refer to DuBuc et al., 2014 here. That paper already showed that U0126 blocks head regeneration.\n8. Lines 289-290 and Fig. 5F. Is the delay in wound healing significant? The sample size is low, so making biological replicates and identifying the confidence interval is important to understand whether there is a delay at all.\n9. Line 295. Can heads re-commence regeneration after U0126 washout? If not, can re-amputation re-start regeneration? Is it possible to create a “dormant Nematostella head” as in https://www.nature.com/articles/s41467-017-02338-x\n10. Lines 325-327. How did the authors come up with 40 embryonic and 40 regeneration genes? Were these the top 40? Please explain.\n11. Lines 327-330. The authors write “From the 40 embryonic GRN genes, 25 have been described as downstream targets of MEK/ERK signalling, while the expression of 15 are controlled by the canonical Wnt (cWnt) pathway at the onset of gastrulation.” Can such a clear distinction really be made? There must be an overlap between the 25 and the 15. For example, WntA and Sp8/9 are clearly not only MEK/ERK but also Wnt/b-catenin targets at gastrulation.\n12. Lines 331-342. I fail to see the message here. These genes are co-expressed, so, indeed, they are possibly doing something together. Please rephrase for more clarity.\n13. Lines 347-349. Johnston et al. write, “Strikingly, 19 out of the 25 identified embryonic MEK/ERK downstream targets (Table S5) are downregulated by U0126 treatments during regeneration (Fig. 6Bi)”. Why is it striking? I would rather say, “as expected”.\n14. Lines 370-372. I do not doubt the involvement of Wnt and MEK/ERK signaling in the regulation of regeneration but I do not see any experimental confirmation of the placement of Wnt downstream of MEK/ERK. This study lacks systematic testing of what happens to regeneration upon upregulation and downregulation of Wnt as well as upregulation and downregulation of MEK/ERK as well as data showing which of the two (or maybe both) pathways is activated by the early wave of apoptosis. Please also do not forget to refer to the Hydra paper of Chera et al., 2009 when talking about the effect of apoptosis on Wnt.\n15. Line 396. Remove “and”\n16. Line 427. Why does the logic have to be reshuffled? What is the basis for this prediction?\n17. Line 429. Remove “and” from “other and vertebrates”.\n18. Lines 460-463. Johnston et al. write that “cWNT and MEK/ERK activate a distinct set of downstream targets, while during regeneration MEK/ERK signalling is able to activate not only embryonic MEK/ERK targets but also embryonic cWNT as well as “regeneration-specific” genes.” This is a “false dichotomy”. There is a clear overlap between cWNT and MEK/ERK targets in the embryo (WntA, Sp8/9 – see Kraus et al., 2016 and Lebedeva et al., 2017). Please correct.\n19. Line 676. Probably the title of the figure should be “ERK phosphorylation is required for regeneration”.\n20. Figure 7A. The authors call this a network module, but it is just a list of MEK/ERK target genes. It will become a network module once the regulatory relationships between the targets are experimentally established.\n21. Figure 7B. This scheme is very suggestive but I do not think there is experimental evidence for it. Has it been shown that b-catenin signaling is required for MEK/ERK signaling in the embryos? I do not think there is a publication showing that b-catenin knockdown leads to the absence of MEK/ERK signaling in the embryos. Similarly, this study did not demonstrate experimentally that MEK/ERK is upstream of Wnt/b-catenin. If this is true, b-catenin target genes should be suppressed upon U0126 treatment, but this has to be checked first.", "reviewer_2": "The manuscript by Warner et al uses a re-analysis of prior transcriptomic data to address the similarities and differences between embryogenesis and regeneration in the model sea anemone Nematostella vectensis. The fundamental question of whether regeneration is simply a redeployment of development has been extensively posed in the literature in prior work. Hence the novelty of this study is primarily restricted to the organismal system and the systems biology methods employed to identify differentially expressed genes. As it appears this part of the work largely draws on analysis of published data, the key contribution amounts to the validation and functional experiments and therefore the impact of the work is fairly field-specific. In that regard, this manuscript provides an interesting starting point but falls significantly below the level of general insight expected for the journal. The experimental validation sections of the paper, which address roles for apoptosis and MEk/ERK signaling in regeneration are a big strength of the paper and represent an important advance in the field.\nOn the whole, the figures presenting the transcriptomic analysis are significantly below journal standards and the data do not clearly support conclusions drawn by the authors. Several highly general statements e.g. “This indicates the transcriptional dynamics of embryogenesis are more profound than those of regeneration” are so vague as to be meaningless as the analysis of bulk RNA-seq data likely obscures the transcriptional dynamics of critical cells at the wound site that contribute to regeneration. In this regard, a much higher resolution understanding could be gained by scRNA-seq or spatial transcriptomic approach. It is not clear whether or how this part of the work will have impact or utility to the broader community beyond serving as a starting point for the subsequent experiments.\nThe ZVAD experiments to inhibit apoptosis reveal an interesting requirement for apoptotic cell death in the initiation of regeneration and reflect one of the stronger logical components of the paper. It would be interesting to add an analysis of cell death following wounding without amputation to determine whether cell death can be seen as a consequence of wounding or reflects a component of the regenerative program proper. The TUNEL analysis suggests there may be wound-site specific and systemic cell death upon amputation and the authors could significantly expand the impact of the work by digging deeper in this area.\nSimilar to the analysis of cell death, the results on MEK/ERK could provide key new insights into Nematostella regeneration. Combined with the work on apoptosis, this is the strongest part of the paper. However, it remains unclear how direct or indirect the activation of downstream targets may be and how specifically U0126 treatment affects MEK/ERK signaling. Given the proliferation of genetic methodologies for Nematostella, are there not more rigorous approaches available to demonstrate specific gene requirements by CRISPR or shRNA knockdown?", "reviewer_3": "The manuscript by Johnston et al. uses a comparative approach to determine the relationship between embryonic development and regeneration at both molecular and cellular levels. While defining the key factors that distinguish regeneration from development is fundamentally important, experimentally, only a few research organisms offer the possibility to address this question. In this context, the authors leveraged the accessibility of embryonic development and regeneration in the sea anemone Nematostella vectensis and have generated and analyzed impressive transcriptomic data covering the temporal dynamics of both embryogenesis and regeneration. Building on this comparative approach and functional data involving ERK signaling, the authors identified and validated gene expression patterns that reflect a rewiring of the embryonic genetic program. At the cellular level, apoptosis appears to be required for regeneration, which emphasizes the role of this cellular process as a core mechanism deployed across different regenerative species. The great strength of this study is that the authors have produced significant transcriptomic resources for the scientific community, and the advancements presented here certainly establish this sea anemone as an attractive research organism to study the developmental origins of regeneration. In general, the experiments are well performed throughout the manuscript. However, the interpretations of some results would require clarification or additional controls to address the following concerns:\nMajor concerns:\n1- The authors claim that “Regeneration is transcriptionally modest compared to embryonic development”. However, this conclusion is based on comparing oral pole regeneration, which does not represent the full regenerative capacity of the polyp while embryogenesis builds a whole organism. The authors should reconsider this statement as the current data do not support it.\n2- An experimental validation of ‘regeneration specific” genes is missing. In this context, an attractive category is the list of genes that are taxonomically restricted. This work has the potential to inform us about their integration in the gene regulatory logic. For instance, Is there any bias in their spatio-temporal deployment during regeneration?\n3- The authors also mentioned that “Apoptosis is specifically required for regeneration” without providing strong evidence to support this claim. Many studies show embryos or larvae might look morphologically normal while cell differentiation can be dramatically impacted when using molecular profiling. In addition, a peak of cell death has been reported during late larval development (Zang and Nakanishi, 2020). An obvious experiment would be to treat late larvae with the pan-caspase inhibitor Z-VAD and test whether development proceeds normally.\n4- It is exciting to see the involvement of apoptosis in Nematostella regeneration. Could the authors explore the transcriptomic data and show the expression of developmental signals that are provided by these dying cells.\nMinor concerns:\n- The labeling of the panels with multiple letters is not simple to follow and creates confusion.\n- There is a typo in “Apoptosis is a and regeneration-specific process.”"}, "author_response": "We thank the reviewer for the relevant comments that lead to a series of additional experiments and the re-organization of our study to better interconnect the transcriptomic aspects with the functional data. We’re convinced that these additional data have helped to significantly improve the manuscript for publication in Nature Communication.\nTo address the connection between apoptosis and MEK/ERK signaling, we have performed additional experiments and show that while apoptosis seems MEK/ERK-independent at 2hpa (Fig. S17), it is dependent of MEK/ERK activity at 20h (Fig. 6 and Fig. S17). Our new data further suggest that the first wave of apoptosis (2hpa) is caspase-dependent (Fig.S8 and Fig. S17) but MEK/ERK independent, while the second wave (20hpa) is caspase-independent (Fig.6b and Fig. S17) but MEK/ERK dependent. This is detailed in the revised discussion and put in relation to recent findings that apoptosis at 20hpa is ROS dependent, while at 2hpa is it not (Vullien et al. 2024). In addition to showing that apoptosis is crucial for regeneration, we now also show that inhibition of apoptosis does not affect wound-healing or the initiation of the tissue-contact between the mesenteries and the body wall epithelia but is required for its maintenance as well as for both, the initiation and maintenance of cell proliferation (Fig. 5).\nFinally, we have performed a transcriptomic analysis following apoptosis that has been used in the context of the study, i.e., comparing it to the transcriptomes obtained after inhibition of MEK/ERK and cWNT signaling, as well as with the genes belonging to embryonic MEK/ERK and cWNT downstream targets or the set of regeneration-specific genes identified in our study (Fig.7,8).\nTo that end, I suggest:\ni) comparing transcriptomes of the ZVAD-treated, U0126-treated and control regenerates at stages 0.5 and 1, identifying the overlap among the regulatory genes significantly affected by the treatments, and characterizing their expression during regeneration;\nWe have performed transcriptomic analyses on 20hpa traited polyps with DMSO (controls), Z-VAD, U0126 or iCRT14 (iCRT14 blocks cWNT signaling) (Fig. 7, S13, S14, and associated supplemental Tables). The overlap between the significantly DOWN or UP-regulated genes shared between all treatments is rather low (5 for the DOWN and 10 for the UP). Their identity and temporal expression pattern that span the entire regeneration process are shown in Fig. 7D. Go term analyses and assessments of the temporal expression of the DOWN- and UP-regulated genes by the various inhibitory treatments confirm the results from the experiments that assessed the relationship between the three pathways (Fig.6), i.e., MEK/ERK controls downstream targets associated to Wnt signaling and apoptosis. Additional information supporting our statement, are indicated in the revised manuscript. Looking globally at the temporal expression patterns (Fig.S14), we observed that MEK/ERK downstream targets seem upregulated during regeneration in three phases, i) in response to injury, ii) at the onset of regeneration (14-36hpa) and iii) during differentiation (96-144hpa). On the other hand, downstream targets of apoptosis and cWNT are mainly activated either shortly after injury or after 36hpa (Fig.S14).\nii) checking whether apoptosis, MEK/ERK signaling, and expression of b-catenin signaling targets are induced by wounds not causing tissue loss (punctures, incisions), and during foot regeneration;\nWe agree that the suggested experiments to compare the amputation/regenerative response with the injury/wound-healing response or aboral regeneration are certainly interesting, however, in our opinion out of scope of the present manuscript. In fact, the present study focuses on the GRN underlying oral regeneration, and its comparison to the GRN underlying embryonic development. We hope that all the additional experiments, that increased our understanding of the main question of the study, nonetheless satisfy the reviewer.\niii) checking the effect of pharmacological upregulation and downregulation of b-catenin signaling and MEK/ERK signaling on apoptosis at the onset of regeneration;\nTo assess the activation pattern of cWNT signaling, we have developed a CRISPR/Cas9 induced KI line by fusing mCherry to the C-terminal of ß-catenin (Fig. 4L, S7) and observed the beginning of ß-cat nuclearization in cells at the amputation site at 12hpa and 24hpa. Using the readouts for the activation of all pathways (pERK for MEK/ERK signaling, TUNEL for apoptosis, and ß-catenin::mCHERRY for cWNT) we have performed additional experiments in line with the focus of our study. In particular, we performed experiments to understand the relationship (i.e., necessity) between the three pathways using inhibitor treatments (Fig. 6). Doing so we were able to show that MEK/ERK signaling is required for apoptosis and cWNT signaling. This is in line with the observation that inhibition of MEK/ERK signaling causes the most severe phenotype by blocking regeneration at stage 0.5 (Fig. 5). While Z-VAD and iCRT14 treatments have no visible effect on pERK activation, our results suggest a mutual regulation between apoptosis and cWNT signaling at the amputation site at 24hpa (Fig. S12).\niv) checking the effect of pharmacological upregulation and downregulation of b-catenin and MEK/ERK on cell proliferation at the onset of regeneration;\nAs our goal is to determine the necessity of the pathways for regeneration, and not an eventual sufficiency (this is another ongoing study in the lab), we have focused this series of experiments only on the downregulation of the pathways. We have assessed the effects of inhibiting MEK/ERK, apoptosis or cWNT on several and complementary readouts specific to the regeneration process such as wound healing, the onset and maintenance of the tissue contact required for the initiation of regeneration, as well as the onset and maintenance of cell proliferation. All the results are shown now in Fig 5. Our results show that all three pathways are required for injury-induced cell proliferation (Fig. 5D,E).\nv) checking the effect of ZVAD on pERK (immunohistochemistry) and expression of b-catenin target genes (ISH) in amputated animals;\nWe have performed pERK staining following Z-VAD treatments at 20hpa for the above-described experiments (Fig. 6) and show that apoptosis is not required for pERK activation. However, the opposite is the case as TUNEL staining is reduced in UO126 conditions (Fig. 6 and Fig.S17). Regarding the expression of b-catenin target genes in amputated animals, we have obtained a list of potential cWNT downstream target genes during regeneration from our transcriptomics analyses (Fig. 7). We assessed and show their temporal expression pattern during regeneration (Fig. S14). ISH of embryonic cWNT downstream targets during regeneration are shown in Fig. 8C.\nvi) attempting to rescue the effect of the ZVAD-mediated inhibition of apoptosis on regeneration by activating b-catenin signaling and/or MEK/ERK signaling.\nWe agree on the interest of these experiments that further address the tissue competence for the impressive regenerative capacity of N. vectenis. They appear to us out of the scope of the current manuscript and are part of another project from the lab. Following the addition of a wealth of new data characterizing the role of these three pathways during regeneration, we have decided to remain focused on the main question of the manuscript – namely, to what extend regeneration recapitulates the embryonic GRN.\nAmong all the proposed treatments, only upregulation of MEK/ERK seems not to have been routinely used in Nematostella. However, since FGFRb is expressed in the oral region of Nematostella, recombinant FGF incubation appears to be a plausible way to achieve upregulation of MEK/ERK.\nIn another study from the lab focusing on the role of the tissue contact and subsequent tissue-crosstalk (Andreoni et al., close to submission), we have a series of evidence suggesting that FGFRb is indeed involved in the injury-induced activation of MEK/ERK and its down-stream targets. However, FGFRb does not control the entire set of downstream targets and other pathways seem to converge in MEK/ERK activation at the onset of regeneration in N. vectenis. Preliminary data for another study from the lab, seems to suggest that recombinant FGF signaling alone is not sufficient to induce regeneration.\nOnce the authors have experimentally analyzed the links and clearly established the regulatory hierarchy between apoptosis, proliferation, MEK/ERK signaling, and Wnt/b-catenin signaling, and identified and characterized the candidate genes responsible for the start of regeneration, the article can be re-considered for publication at Nature Communications.\nWe thank the reviewer again for the constructive comments. We hope that the additional data sets and detailed analyses of the three pathways during regeneration provide compelling evidence that MEK/ERK signaling is the earliest and most crucial injury-induce signal that triggers an apoptosis and cWNT dependent regenerative response.\nBased on our initial data and the new ones we acquired during the revisions, we propose that all three pathways are required for regeneration, the initiation and maintenance of cell proliferation. We further show that MEK/ERK signaling has the most drastic effect in launching the regeneration process (Fig. 5). In line with this idea, we show that MEK/ERK signaling is required for both apoptosis and cWNT signaling at the onset of regeneration (20hpa). Furthermore, our results, in particular the data from Fig. 6 (that specifically addresses the relationships) and the RNAseq analysis (Fig. 7) also support more subtle and non-hierarchical relationships, i.e., the mutual control of apoptosis and cWNT signaling. Importantly, our results enable us to perform a compelling comparison between the deployment of these pathways during embryonic development and regeneration highlighting a reshuffled hierarchy (Fig. 9).\nIn addition to the general suggestions above, I list specific things for the authors to address in the revision (in order of appearance in the text, not in order of importance).\n1. Lines 61-62. Johnston et al. write about genes “specifically expressed or required during regeneration” as opposed to “genes specific to embryonic development” and “re-used” genes, and refer to a paper showing the involvement of Sox2 in the regeneration but not in the development of the hair follicle. This is misleading – Sox2 is regeneration-specific only in the hair follicle context, but it is a crucial regulator of the embryonic development.\nWe have added “initial development” to the manuscript.\n2. Lines 93-98. Johnston et al. certainly addressed i) and ii). I am not sure about how the “plasticity of the network architecture” (iii) was demonstrated, and I do not think that the paper shows anything with regard to the “rewiring” (iv). If Fig. 7B is meant, I do not find that this paper convincingly proves experimentally that MEK/ERK is upstream of Wnt/b-catenin signaling during regeneration. If the authors refer to lines 344-362, I am not sure I see evidence for rewiring there: most of the embryonic MEK/ERK targets are still MEK/ERK targets during regeneration.\nWe are convinced that the additional sets of experiments we performed for the revision of this manuscript have largely addressed the concerns raised by the reviewer, in particular that MEK/ERK is upstream of cWNT signaling. The term “rewiring” is mainly linked to two observations we made: A) the simplified temporal deployment of cWNT -> MEK/ERK -> Apoptosis during embryonic development is not conserved during regeneration but reshuffled with the following temporal deployment of MEK/ERK -> Apoptosis -> cWNT during regeneration. B) During embryonic development, cWNT activates a largely distinct set of downstream targets than MEK/ERK; During regeneration, these distinct set of downstream targets are controlled by MEK/ERK. More specifically, while most (not all) embryonic MEK/ERK targets are still MEK/ERK downstream targets during regeneration, a subset of embryonic cWNT downstream targets is controlled by MEK/ERK but not anymore by cWNT during regeneration (Fig. 8A). Thus, and because “regeneration-specific” genes are also controlled by MEK/ERK signaling during regeneration, we propose that regeneration redeploys elements of the embryonic gene regulatory network but in a different temporal sequence and rewired / interconnected with “regeneration-specific” genes.\n3. Lines 117-122. How exactly are the phases defined? If by transcriptomic changes, there probably should be some metric to it. How do they correspond to the morphological phases of Amiel et al., 2015?\nTo clarify this point, we added supplementary information to this section.\n4. Lines 152-154. How exactly do the authors define “regeneration-specific genes”? I did not check all the 124 genes, of course, but I am sure one cannot call Wntless a “regeneration-specific gene”. According to the authors’ own NvERTx database, Wntless is a maternally deposited transcript, and Wntless gene is strongly expressed throughout the development, which is absolutely not surprising, given the importance of Wnt signaling in development and the fact that Wntless is a critical member of the Wnt secretion pathway.\nThe reviewer is correct and that is why we defined the “regeneration-specific” genes not because they are only expressed during regeneration or that their roles are only associated to regeneration, but that their expression is dynamic during regeneration (fC > 2-fold) and not during embryonic development (fC < 2-fold). While we have amended the first phrase of this section to make this clearer, the text referring to Figure 1F stated, and still states, that a part of the “regeneration-specific” genes is expressed during embryonic development.\n5. Lines 241-243. The finding of the requirement of apoptosis for regeneration but not for normal development is among the most interesting things reported in the paper. Currently, the results of TUNEL on embryos are “data not shown”. Please show and quantify these data and add them to Fig. 4Bii!\nWe have added the experiment that shows that apoptosis is required for regeneration and not for embryonic development to Fig. 5A. We now also cite the manuscript that showed that TUNEL positive cells appear in the larvae during metamorphosis (Zang & Nakanishi, 2020).\nPlease provide quantification data (Fig. 4Bii) in a form of box plots or individual datum points rather than a table with means, so that it is possible to estimate the variance between polyps. I also suggest increasing N to at least 5 in every case.\nDone.\n6. Lines 259-260. The authors suggest that apoptosis induces cell proliferation. In order to make the argument that apoptosis is upstream of proliferation more convincing, I suggest directly blocking proliferation (with cyclin-dependent kinase or DNA polymerase inhibitors) and checking whether apoptosis is still detectable.\nWhen blocking directly proliferation using Hydroxy Urea, regeneration is blocked at stage 1 (Amiel at al. 2015). However, we do not understand how directly blocking proliferation will inform us if apoptosis is upstream of cell proliferation. From our understanding, such experiment will only suggest a potential impact of cell proliferation on apoptosis and not the contrary. By using the pan-caspase inhibitor Z-VAD, we showed that the inhibition of apoptosis blocks not only the initiation but also the maintenance of injury-induced cell proliferation during regeneration (Fig.5D). This shows that apoptosis is upstream to and required for cell proliferation.\n7. Line 270. The authors have to refer to DuBuc et al., 2014 here. That paper already showed that U0126 blocks head regeneration.\nWe have added Dubuc et al 2014 (that we cite elsewhere multiple times and that mainly focused on wound-healing) to the introduction, mentioning that this study suggests that MEK/ERK signaling is required for oral regeneration (without showing the specific numbers that support their observation).\n8. Lines 289-290 and Fig. 5F. Is the delay in wound healing significant? The sample size is low, so making biological replicates and identifying the confidence interval is important to understand whether there is a delay at all.\nThe wound-healing assay on U0126 treated regenerating polyps has been performed in replicates that have been pooled/integrated in Figure 5B (indicating the number of assessed polyps). This figure also shows our assessment of the effects of blocking apoptosis and cWNT signaling on wound healing. Statistical analyses confirmed the delay in wound healing for U0126 treated polyps, while iCRT14 and Z-VAD slightly accelerated the process.\n9. Line 295. Can heads re-commence regeneration after U0126 washout? If not, can re-amputation re-start regeneration? Is it possible to create a “dormant Nematostella head” as in https://www.nature.com/articles/s41467-017-02338-x\nWhile this is indeed interesting, we consider these experiments out of the scope of the present manuscript and are part of another ongoing project in the laboratory.\n10. Lines 325-327. How did the authors come up with 40 embryonic and 40 regeneration genes? Were these the top 40? Please explain.\nWe have rephrased this section in the manuscript and hope this is clearer now.\nTo identify the 40 embryonic and the 40 regeneration genes, we have taken the list of the embryonic i) MEK/ERK or ii) cWNT downstream targets identified in Röttinger et al, 2012 and Amiel et al. 2017 respectively, as well as iii) the “regeneration-specific genes” we identified in this current study. Then, we have cross-referenced these three gene lists with the differentially expressed genes at 20hpa. Doing so we have identified the mentioned 40 genes associated to embryonic MEK/ERK and cWnt downstream targets as well as 44 “regeneration-specific” genes (we updated this number in the manuscript, Fig. 3A,B; S3).\n11. Lines 327-330. The authors write “From the 40 embryonic GRN genes, 25 have been described as downstream targets of MEK/ERK , while the expression of 15 are controlled by the canonical Wnt (cWnt) pathway at the onset of gastrulation.” Can such a clear distinction really be made? There must be an overlap between the 25 and the 15. For example, WntA and Sp8/9 are clearly not only MEK/ERK but also Wnt/b-catenin targets at gastrulation.\nBased on the data from the cited papers (Röttinger et al 2012 and Amiel et al, 2017) our statement is largely correct. Figure 7 from Amiel et al, 2017 shows the downstream targets of MEK/ERK and cWnt signaling during the onset of gastrulation. We have attentively compared the pathway specific downstream targets (Table S5, S6) and the only genes that indicate an input from cWnt and MEK/ERK during embryonic development are nfix-like, hd050, elk-like1 and gsc. If the reviewer can send us the references that show that wntA and sp8/9 are both controlled by MEK/ERK and cWnt at the onset of gastrulation, we would be happy to include these references.\nWe have re-written this section and to consider that not strictly all genes are downstream of one or the other pathway, we reformulated the sentence in the rewritten paragraph. “These embryonic downstream targets are largely pathway specific as only 4 genes (nfix-like, hd050, elk-like1 and gsc) are controlled by MEK/ERK and cWNT at the onset of gastrulation (Table S5,6).”\n12. Lines 331-342. I fail to see the message here. These genes are co-expressed, so, indeed, they are possibly doing something together. Please rephrase for more clarity.\nWe make a distinction between “co-expression” and “syn-expression”. Co-expression can be temporal (i.e., the expression patterns shown in Figure S2) or spatial (i.e., some of the expression pattern shown in Figure 5). Syn-expression, as defined by Niehrs et al., 2000 (that we now cite for additional information) are genes that are expressed identically at both level spatial and temporal. In short, genes that belong to syn-expression groups are more likely to participate in the GRN underlying a common biological process, compared to genes that share either temporal or spatial co-expression.\n13. Lines 347-349. Johnston et al. write, “Strikingly, 19 out of the 25 identified embryonic MEK/ERK downstream targets (Table S5) are downregulated by U0126 treatments during regeneration (Fig. 6Bi)”. Why is it striking? I would rather say, “as expected”.\nTaking into account the reviewer’s comment and the new data, we have removed the word “strikingly” from the sentence. The reason we used “strikingly” was that considering the different context between embryonic development and regeneration, we found this observation striking and not necessarily expected. Additional data, particularly the observation that inhibiting cWnt signaling during regeneration does not affect embryonic cWnt downstream targets (Fig.8), confirm our idea that conservation of 'pathway-specific' downstream targets is not necessarily expected.\n14. Lines 370-372. I do not doubt the involvement of Wnt and MEK/ERK signaling in the regulation of regeneration but I do not see any experimental confirmation of the placement of Wnt downstream of MEK/ERK. This study lacks systematic testing of what happens to regeneration upon upregulation and downregulation of Wnt as well as upregulation and downregulation of MEK/ERK as well as data showing which of the two (or maybe both) pathways is activated by the early wave of apoptosis. Please also do not forget to refer to the Hydra paper of Chera et al., 2009 when talking about the effect of apoptosis on Wnt.\nOur additional experiments for the revised version of the manuscript should satisfy the reviewers’ comments. The Chera et al. paper was cited and discussed in the initial version of the manuscript (l. 441, l.449) and in the revised version in l.624, l.676, l.691.\n15. Line 396. Remove “and”\nDone.\n16. Line 427. Why does the logic have to be reshuffled? What is the basis for this prediction?\nSee reply to the 2nd major comment above.\n17. Line 429. Remove “and” from “other and vertebrates”.\nDone.\n18. Lines 460-463. Johnston et al. write that “cWNT and MEK/ERK activate a distinct set of downstream targets, while during regeneration MEK/ERK signalling is able to activate not only embryonic MEK/ERK targets but also embryonic cWNT as well as “regeneration-specific” genes.” This is a “false dichotomy”. There is a clear overlap between cWNT and MEK/ERK targets in the embryo (WntA, Sp8/9 – see Kraus et al., 2016 and Lebedeva et al., 2017). Please correct.\nWe’re unclear, or even in disagreement with the reviewers’ interpretation from the cited References in regard to our study.\nIn Kraus et al. 2016, the study nicely describes a pre-bilaterian blastoporal axial organizer. We could not find any sign of the gene SP8/9, any inhibition of MEK/ERK, or any other experiment that would suggest a control of wntA expression by MEK/ERK. The Kraus et al. 2016 study mainly used the cWnt activator 1-Azakenpaullone, and showed that among the tested genes, wntA can respond to 1-Azakenpaullone at the gastrula stage. These data extended previous observations from Röttinger et al, 2012 to later stages (blastula versus gastrula, respectively). It is important to note here that in Kraus et al. 2016, the experiments correspond to global overactivation of the cWnt pathway that only show that wntA can respond to this global overactivation. Meaning that this is no evidence for wntA being a downstream target of cWnt signaling in normal developmental conditions. In fact, when performing Knock-Down experiments using a morpholino targeting TCF, the effector of cWnt signaling, wntA expression seems not affected, suggesting that wntA is not a target of cWNT signalling in this early developmental context (Fig. 10 in Röttinger et al, 2012). In line with these data, injection of a morpholino against ß-catenin does not block wntA expression neither, but instead, extend its expression territory to more aboral tissues, suggesting that cWnt signaling is involved in restricting expression of wntA to the oral central ring domain (Leclere et al. 2016). Taken together, these data indicate that during embryogenesis wntA is NOT a downstream target of cWnt signaling at the oral pole under physiological developmental conditions at the late blastula stage in Nematostella.\nOn the other hand, wntA expression was downregulated under U0126 (MEK inhibitor) conditions at the late blastula (Layden et al, 2016; Additional File 8 – Table S8). Although, the injection of a morpholino targeting erg, a potential effector of MEK/ERK signaling during embryonic development of Nematostella (Amiel et al, 2017), did not yield in a significant downregulation of wntA expression. These data indicate that wntA is an Erg-independent MEK/ERK downstream target.\nThe gene sp6/8 has also been identified in the microarray performed following UO126 treatments in late blastula stages (Layden et al, 2016; Additional File 8 – Table S8). Like wntA, sp6/8 is not downregulated following the injection of morpholinos targeting erg (Amiel et al, 2017) showing that sp6/8 is also an Erg-independent MEK/ERK downstream target.\nConcerning the second reference indicated by the reviewer, Lebedeva et al., 2017, we could not find any publication corresponding to Lebedeva et al, 2017. In fact, the Google Scholar from Tatiana Lebedeva (which we believe is the author the reviewer is referring to) indicates only one paper for 2017, however, under the name of Tatiana Bagaeva. This paper addressed a Nematostella un-related topic.\nThe closest publication regarding the reviewers comment we could find and in which Tatiana Lebedeva is 1rst author corresponds to Lebedeva et al, 2021 (DOI: 10.1038/s41467-021-24346-8). This publication includes a sp gene, but it is called sp6-9, and not sp8/9 (which corresponds to the reviewer’s concern). Although the spatial expression patterns are similar between the gene sp6/9 (Lebedeva et al, 2021) and the gene sp6/8 (from Layden et al 2016 and Amiel et al, 2017, which could be the gene that has been named sp6/9 by Lebedeva et al, 2021), we could not find any sequence related to sp6-9 to confirm this similarity.\nIn Lebedeva et al. 2021, experiments have been carried out at the gastrula stage and not at the blastula stage like in Röttinger et al 2012; Layden et al 2016 or Amiel et al 2017. Lebedeva et al. 2021 has identified sp6-9 in a transcriptomic analysis following 1-Azakenpaullone treatments (meaning in a cWnt overactivation context) showing that sp6-9 can respond to global cWnt overactivation. In another set of experiments sp6-9 expression has been assessed by in situ hybridization following Knock Down of genes of interests (bra, lmx, foxA or foxB) or/and 1-Azakenpaullone treatments. These experiments shows that sp6-9 expression is shifted orally when the cWnt downstream targets bra, lmx, foxA or foxB (all four expressed in the central ring) are functionally impaired. These results highlight the importance of these four genes (and cWnt signalling) for spatial restriction, confining sp6-9 expression to the external ring and thus patterning the O-A axis during gastrulation. The 1-Azakenpaullone treatments under these KD conditions further confirm that sp6-9 expression can be activated in global cWnt overexpression conditions, independently to the absence of one of those genes. In sum, we could not find any results that suggest that sp6-9 expression is controlled by cWnt under physiological developmental conditions at the late blastula stage in Nematostella.\nTaken together, we do not see how the mentioned publications from the reviewer show that the data we used for the comparison leading to our conclusions is a “false dichotomy”. As mentioned above we have slightly tempered our statement to take into account the genes, for which we have found evidence that they are controlled by MEK/ERK and cWNT signaling during embryonic development.\n19. Line 676. Probably the title of the figure should be “ERK phosphorylation is required for regeneration”.\nWe agree with the reviewer and changed the title.\n20. Figure 7A. The authors call this a network module, but it is just a list of MEK/ERK target genes. It will become a network module once the regulatory relationships between the targets are experimentally established.\nWe agree and have removed that sentence from the legend.\n21. Figure 7B. This scheme is very suggestive but I do not think there is experimental evidence for it. Has it been shown that b-catenin signaling is required for MEK/ERK signaling in the embryos? I do not think there is a publication showing that b-catenin knockdown leads to the absence of MEK/ERK signaling in the embryos. Similarly, this study did not demonstrate experimentally that MEK/ERK is upstream of Wnt/b-catenin. If this is true, b-catenin target genes should be suppressed upon U0126 treatment, but this has to be checked first.\nWe agree with the reviewer. It was meant to indicate the temporal relationship rather than functional relationship. The new version of the figure (now Fig. 9) now focuses on the temporal deployment of the studied pathways rather than the functional relationships.\nReviewer #2 (Remarks to the Author):\nThe manuscript by Warner et al uses a re-analysis of prior transcriptomic data to address the similarities and differences between embryogenesis and regeneration in the model sea anemone Nematostella vectensis. The fundamental question of whether regeneration is simply a redeployment of development has been extensively posed in the literature in prior work. Hence the novelty of this study is primarily restricted to the organismal system and the systems biology methods employed to identify differentially expressed genes. As it appears this part of the work largely draws on analysis of published data, the key contribution amounts to the validation and functional experiments and therefore the impact of the work is fairly field-specific. In that regard, this manuscript provides an interesting starting point but falls significantly below the level of general insight expected for the journal. The experimental validation sections of the paper, which address roles for apoptosis and MEk/ERK signaling in regeneration are a big strength of the paper and represent an important advance in the field.\nWe thank the reviewer for the comments and hope that the revised version of the manuscript provides more detail and experimental evidence to support our claims. Previous studies that have addressed the question were and are presented in the manuscript and we strongly believe that our study goes further than what has been described so far. To address the reviewer’s comments, we have conducted a significant number of additional experiments and functional analyses for this revised manuscript, which we believe will further increase the interest within the broader scientific community.\nOn the whole, the figures presenting the transcriptomic analysis are significantly below journal standards and the data do not clearly support conclusions drawn by the authors. Several highly general statements e.g. “This indicates the transcriptional dynamics of embryogenesis are more profound than those of regeneration” are so vague as to be meaningless as the analysis of bulk RNA-seq data likely obscures the transcriptional dynamics of critical cells at the wound site that contribute to regeneration.\nWe have added a sentence to the results section to place this sentence better into the comparative context of the manuscript. As the amputation we performed were below the pharynx (~1/3-1/2 of the body of juveniles), the program that is deployed during regeneration must allow the reformation of the main parts and structures of the animal (i.e., body-wall epithelia, mesenteries, pharynx, mouth, tentacles as well as the associated muscles and neural networks).\nAs the reviewer noted, the series of transcriptomics analyses presented in Fig 1 and 2 (and associated supplemental figures S1, S2, S3) were the starting point for a series of experiments related to the global question we addressed. They are a crucial aspect of our question we’re addressing, i.e., to what extend regeneration re-deploys embryonic gene networks. Thus, we decided to keep that statement as it relates not only to Fig. 1D but also to Fig. 1E and Fig. 3 leading to the global conclusion that regeneration is largely a partial re-use of embryonic development.\nIn this regard, a much higher resolution understanding could be gained by scRNA-seq or spatial transcriptomic approach. It is not clear whether or how this part of the work will have impact or utility to the broader community beyond serving as a starting point for the subsequent experiments.\nThe transcriptomic data we used and completed for our study (Warner et al., 2018) is already used by the community for various studies covering development and regeneration in a variety of research models (80 citations as of today). For the current study, we have chosen to use these bulk RNAseq data on purpose as we wanted to address the question on the relationship between embryonic development and regeneration as large as possible and as unbiased as possible, considering also the tissular aspects (i.e. the importance of the mesenteries to induce regeneration – Amiel et al – BioRxiv). Doing so we were able to span a large set of time points during embryonic development (22 time points) and regeneration (16 time points) enabling us to perform an extensive comparison between these two developmental trajectories. The outcome of the first part serves indeed as the starting point for the subsequent experiments of the current manuscript, but also open various venues for additional studies within the lab and the broader community.\nWe agree with the reviewer that the cell specific dynamics and regulatory sc- or spatial RNAseq would yield interesting complementary data. While we consider that these approaches will address other more specific questions (ex the comparison of cell fate trajectories), it is currently out of the scope for the present study and part of ongoing work in the laboratory.\nThe ZVAD experiments to inhibit apoptosis reveal an interesting requirement for apoptotic cell death in the initiation of regeneration and reflect one of the stronger logical components of the paper. It would be interesting to add an analysis of cell death following wounding without amputation to determine whether cell death can be seen as a consequence of wounding or reflects a component of the regenerative program proper.\nIn both conditions, puncture/wounding (non-regenerative – Dubuc et al. 2014) or amputation (regenerative – our study), the first wave of apoptosis is a consequence of wounding. The results from Dubuc et al. 2014 were and are still mentioned in our manuscript. However, the second and third waves of apoptosis we have identified and that are not located at the wound site anymore, seem to reflect a component of the regenerative program.\nThe TUNEL analysis suggests there may be wound-site specific and systemic cell death upon amputation and the authors could significantly expand the impact of the work by digging deeper in this area.\nWe agree with the reviewer that this is an interesting line of research and have performed additional experiments to gain a better understanding of the different waves of apoptosis (Fig.S9, briefly described above). We also highlight and discuss that the first wave of apoptosis is caspase-dependent but ERK-independent, while the 2nd wave of apoptosis seems caspase-independent, while ERK-dependent.\nSimilar to the analysis of cell death, the results on MEK/ERK could provide key new insights into Nematostella regeneration. Combined with the work on apoptosis, this is the strongest part of the paper. However, it remains unclear how direct or indirect the activation of downstream targets may be and how specifically U0126 treatment affects MEK/ERK signaling. Given the proliferation of genetic methodologies for Nematostella, are there not more rigorous approaches available to demonstrate specific gene requirements by CRISPR or shRNA knockdown?\nWe agree with the reviewer on the various points that were raised. ATACseq to gain insight into the regulatory elements and determine direct or indirect interactions during regeneration is part of another ongoing study from the laboratory and we decided to keep this aspect out of the present manuscript. We did this also for the sake of maintaining the same level of comparison between embryonic development and regeneration. As for the use of other functional approaches, unfortunately, we and the community are not quite there yet. Currently, there are no available conditional KO approaches, and our attempts to soak morpholino or siRNA in juveniles did not produce exploitable results.\nReviewer #3 (Remarks to the Author):\nThe manuscript by Johnston et al. uses a comparative approach to determine the relationship between embryonic development and regeneration at both molecular and cellular levels. While defining the key factors that distinguish regeneration from development is fundamentally important, experimentally, only a few research organisms offer the possibility to address this question. In this context, the authors leveraged the accessibility of embryonic development and regeneration in the sea anemone Nematostella vectensis and have generated and analyzed impressive transcriptomic data covering the temporal dynamics of both embryogenesis and regeneration. Building on this comparative approach and functional data involving ERK signaling, the authors identified and validated gene expression patterns that reflect a rewiring of the embryonic genetic program. At the cellular level, apoptosis appears to be required for regeneration, which emphasizes the role of this cellular process as a core mechanism deployed across different regenerative species. The great strength of this study is that the authors have produced significant transcriptomic resources for the scientific community, and the advancements presented here certainly establish this sea anemone as an attractive research organism to study the developmental origins of regeneration. In general, the experiments are well performed throughout the manuscript. However, the interpretations of some results would require clarification or additional controls to address the following concerns:\nMajor concerns:\n1- The authors claim that “Regeneration is transcriptionally modest compared to embryonic development”. However, this conclusion is based on comparing oral pole regeneration, which does not represent the full regenerative capacity of the polyp while embryogenesis builds a whole organism. The authors should reconsider this statement as the current data do not support it.\nWe thank the reviewer to point this out and have added “oral” and replaced “is” by “appears” in this sentence and included the raised comment in the manuscript. Rather than a contradiction, we consider the experimental design for the comparison we carried out (embryonic development vs oral regeneration) an explanation for the observed difference in transcriptional dynamics. We have included this in the updated discussion that was also extended to explain possible differences with other studies.\nThe body parts following amputation that are used for the transcriptomic analysis cover the reformation of a large part of the body. In fact, the sub-pharyngeal amputation we performed on the juveniles, measuring ~500µm in length, are nearly corresponding to mid-body amputation. This means that the molecular program deployed during regeneration must allow the reformation of most of the body of the animal (i.e., body-wall epithelia, mesenteries, pharynx, mouth, tentacles as well as the associated muscles and neural networks).\nRegeneration in Nematostella occurs only when mesenterial tissues are present (Amiel et al, BioRxiv), making it impossible to launch regeneration from a mesentery-less physa. To cover the largest possible regenerative program, we have decided to perform the transcriptome on entire regenerating polyps and not only the amputation site. Furthermore, systemic gene expression changes in tissues distant from the site of injury has been documented in numerous model systems, including in Nematostella as shown in a recent study (Cheung et al, 2025). Expression patterns induced in the mesenteries after amputation (ex. Fig. 8C) are in line with this idea. Thus, our data and results consider any patterning and scaling event even far from the amputation site and making our intra-species comparison as broad and as unbiased as possible.\n2- An experimental validation of ‘regeneration specific” genes is missing. In this context, an attractive category is the list of genes that are taxonomically restricted. This work has the potential to inform us about their integration in the gene regulatory logic. For instance, Is there any bias in their spatio-temporal deployment during regeneration?\nA first, global, validation of the regeneration-specific genes set, that include the pro-apoptotic gene bax, is the assessment of the role of apoptosis. This revealed that compared to embryonic development (and taking into account the reference mentioned by the reviewer below), apoptosis seems to be a regeneration-specific process. As the focus of the present study is to understand to what extend regeneration redeploys embryonic elements, we have included in situ hybridizations for regeneration-specific genes in Fig. 3 (screen of embryonic MEK/ERK or cWnt downstream targets as well as the regeneration-specific genes up-regulated at 20hpa) and 8 (regeneration-specific genes down-regulated by U0126 treatments). An in-depth functional assessment of the role of these genes during regeneration is part of an ongoing project in the laboratory.\nConsidering the reviewer’s comment and to provide additional insight into the expression of the regeneration specific genes, we have added a table indicating the regeneration expression clusters these genes belong to (Fig. 1F), highlighting an increased number of regeneration specific genes that are part of the expression cluster R9 (36 genes) and R6 (29 genes), followed by R2 (14 genes) and R3 (13 genes).\nMore globally, the manuscript, presents spatial expression patterns for 5 regeneration specific genes: bax, wls, carm, cad & egln (Fig. 3 & Fig. S4; Fig. 4, Fig. 8). All genes belong to expression cluster R6, however, their expression pattern is not identical within the various tissues at the amputation site (i.e., body wall epithelia vs mesenteries), suggesting complementary regulatory elements that drive their expression following amputation. Deciphering this in detail as well as the role of these genes during the process of regeneration is out of scope for the current manuscript and is currently being investigated in another project from the lab.\n3- The authors also mentioned that “Apoptosis is specifically required for regeneration” without providing strong evidence to support this claim. Many studies show embryos or larvae might look morphologically normal while cell differentiation can be dramatically impacted when using molecular profiling. In addition, a peak of cell death has been reported during late larval development (Zang and Nakanishi, 2020). An obvious experiment would be to treat late larvae with the pan-caspase inhibitor Z-VAD and test whether development proceeds normally.\nWe thank the reviewer for pointing out the publication from Zang & Nakanishi that we have missed. This reference is now cited in our manuscript and the statement adjusted accordingly. We have now included experiments (Figure 4A) that show not only that embryonic development occurs normally, but also that metamorphosis occurs following Z-VAD treatments (0-polyp & planula-polyp) at the same rate than DMSO treated embryos.\n4- It is exciting to see the involvement of apoptosis in Nematostella regeneration. Could the authors explore the transcriptomic data and show the expression of developmental signals that are provided by these dying cells.\nWe agree that this is an interesting result. Although the more in-depth analysis of the developmental signals provided by apoptotic cells appear to be out of the scope of the present paper, we performed additional transcriptome for the current study including under apoptosis inhibition during regeneration. The transcriptomic data following Z-VAD treatments indicates a series of developmental genes such as “FoxL, Nidogen, Neuropilin, Bmp1, etc. ” (Fig. 7D, Table SX). While from the current data it seems unlikely that Wnt’s, like in Hydra or Zebrafish, are signals emitted by apoptotic cells, our results could indicate that apoptosis participates in modulating the extracellular matrix and thus the associated signaling.\nMinor concerns:\n- The labeling of the panels with multiple letters is not simple to follow and creates confusion.\nWe have revisited the Figure labelling and hope that the new version facilitates the lecture.\n- There is a typo in “Apoptosis is a and regeneration-specific process.”\nDone."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have significantly improved the paper, and experimentally it is now a very solid work. There are several specific things, which, in my opinion, need to be done for the manuscript to become publishable (see the point-by-point below). I also suggest that the authors shorten the discussion making it less speculative and focusing it on the implications of their main findings rather than on what other questions would be potentially interesting to address.\nPoint-by-point comments in order of appearance in the text (not in the order of importance!).\nLine 163. “Regeneration-specific” genes may be regeneration-specific only in this particular comparison. For example, wntless is a maternal transcript abundantly present throughout development in the pharyngeal and body wall ectoderm.\nLine 333. Fig. S8: In contrast to the staining on Fig. 4L, where nuclear b-cat-mCherry is certainly real at 12 and 24 hpa, the authors misinterpret the b-cat-mCherry staining on Fig. S8D (by the way, D is present in the legend but missing on the figure!). The small circles in the middle of the cells, which the arrowheads are pointing at are not nuclei. They are crowns of microvilli surrounding the cilium, - a similar picture is shown in Fig. 1 of the Momose et al., 2012 on Clytia, and on your own Fig 4L in the 48 hpa image. Nuclei are much bigger - see DAPI. Also, the authors show significant depletion of the mCherry signal in the cell contacts in iCRT14 in comparison with DMSO, which should not happen, since iCRT14 prevents b-cat/Tcf interaction, not something else.\nAnother point: I am not sure what “a”, “b”, “abc”, “ac” and “d” mean on top of the error bars on (C), but to me this plot shows that AZ is the only chemical with a clear effect on b-cat signaling. iCRT14 does not have a significant effect on b-cat in comparison to DMSO. It is not clear to me, how iCRT14 can abrogate the effect of AZ if it does not have an effect on topflash-driven expression when administered alone. A good control for the functionality of iCRT14 would be qPCR or RNA-Seq on b-cat target genes during regeneration. It should be reciprocal in iCRT14 and AZ - at least for genes upregulated in AZ.\nLines 334-335: Clumsy sentence. MEK/ERK and cWnt do not block development and patterning. Experimental modulation of MEK/ERK and cWnt does.\nLines 352-353: The authors write “iCRT14 treated polyps did not display ß-catenin nuclearization at 24hpa (Fig.S8C)” – but their controls also do not show b-cat nuclearization - at least on Fig. S8. See comment to Line 333. There is clear nuclear b-cat-mCherry signal on Fig. 4L at 12 and 24hpa.\nLine 363: Replace “as” with “in comparison to”\nLine 445: On Fig. 7C, the GO-terms are completely unreadable without a 5x zoom, and certainly not on the printout. If this is a main text figure, every part of it has to be readable on a standard page without zooming.\nLines 451-461: How do the authors explain that MAPK signaling is upstream of Wnt, but the genes controlled by Wnt are not affected by MAPK inhibition?\nLines 476-481: The authors write “Systematically comparing these pools of genes (emb_cWnt, emb_ERK, regen_spe) with the up- or down-regulated genes following inhibition of MEK/ERK (Tables S20,S21), apoptosis (Tables S22,S23), or cWnt (Tables S24,S25) during regeneration, we observed that MEK/ERK signaling during regeneration has the most widely extended control on gene expression for all three pools (Fig.8; Fig.S15).” This makes total sense, but doesn’t it contradict what they wrote on lines 451-461 (that target lists barely overlap)?\nLines 495-502: I do not find labels like Bi, Bii, and Biii on Fig. 8. I also do not see any insets on Fig. 7B.\nLine 514: This is not a GRN. Unlike anything published by Eric Davidson, who was using the same type of graphics for GRN representation, here, it is just a list of downstream targets of MEK/ERK without any experimentally proven or even hypothetical regulatory links between them. It occupies figure space without providing any information a supplementary table cannot provide. Please remove.\nLine 534-536: The authors write that their results “support a model in which regeneration reuses core developmental programs such as cell type specification and patterning, while also activating unique, regeneration-specific regulatory elements and transcription factors”. They may be right in principle, but their results do not show anything of this sort. Out of 123 “regeneration-specific genes” in Table S1, there is a single transcription factor, Msx2-like, however, single-cell transcriptomics atlas of Cole et al., 2024 shows that it is strongly expressed in the retractor muscle and also in one particular neuroglandular cell type in a normal adult polyp. So, it may be “regeneration-specific” only in the comparison made by Andreoni-Pham et al., but not otherwise. Since there is no ATAC-Seq data in the manuscript, the authors cannot say that their data supports the existence of “regeneration-specific regulatory elements” either.\nLine 657: wnt7\nLine 703: The scheme on Fig. 9 is incorrect. MEK/ERK is responsible for the initiation of the endomesodermal expression in the plate in the 6-8 hpf blastula stage (Haillot et al., 2025); that is the same time when maternal beta-catenin goes into the nuclei of all future ectodermal cells (Lebedeva et al., 2025). So, there has to be not only the orange but also the green arrows to each embryo cartoon.", "reviewer_2": "The authors have incorporated significant new functional experiments into the manuscript in F5 and F6. This addresses one major concern. Still, their rebuttal to my comments offered mostly argumentation and minimal specific changes in response to the original comments (excepting one changed sentence and the analysis of the waves of apoptosis in SF9). On the whole, the paper is improved and scientifically sound but I maintain some concerns about the generality of the bulk RNA-seq analysis, the lack of single cell resolution, and thus the general impact of the work relative to the current literature.", "reviewer_3": "The revised manuscript by Andreoni-Pham, Johnston & Warner et al. represents a substantial improvement over the initial submission. The authors have effectively addressed the major concerns by incorporating additional experiments that strengthen the connections between their transcriptomic analyses and claims. The manuscript now presents a more cohesive narrative that integrates descriptive and functional approaches. I believe the work is now suitable for publication in Nature Communications."}, "author_response": "We thank the reviewer for highlighting the improvements of our revised manuscript as well as for the additional constructive point-by-point comments / recommendations that we addressed below and in the manuscript. We have also revisited and shortened the discussion.\nPoint-by-point comments in order of appearance in the text (not in the order of importance!).\nLine 163. “Regeneration-specific” genes may be regeneration-specific only in this particular comparison. For example, wntless is a maternal transcript abundantly present throughout development in the pharyngeal and body wall ectoderm.\nWe agree that wntless is not solely expressed during regeneration but also during embryonic development. In fact, as stated on l. 160-161 from the 123 “regeneration-specific” genes, “..78 genes are expressed during embryonic development but are not considered dynamic from our differential expression analysis above (Fig.1F)”. wntless is among these 78 genes.\nAs stated in the text (see l.154-157) and in our previous reply to a similar comment, we defined the “regeneration-specific” genes not because they are only expressed during regeneration, but that their transcriptional expression is dynamic during regeneration (fC > 2-fold) and not during embryonic development (fC < 2-fold). As stated in l.158-159, from the 123 “regeneration-specific” genes highlighted in this analysis, only “45 are detectable specifically during the regeneration process indicating they are transcriptionally silent during embryogenesis”. We have replaced “specifically” by “exclusively” to clarify this point.\nAlthough we consider that we have clearly defined the term “regeneration-specific”, we remain cautiously open to replacing this concise term into a longer description. Specifically, the term “regeneration-specific” could be replaced by “genes with a regeneration-specific expression dynamics”. But as we use the term frequently throughout the manuscript, this would make it more heavy to read and extend the manuscript length.\nLine 333. Fig. S8: In contrast to the staining on Fig. 4L, where nuclear b-cat-mCherry is certainly real at 12 and 24 hpa, the authors misinterpret the b-cat-mCherry staining on Fig. S8D (by the way, D is present in the legend but missing on the figure!). The small circles in the middle of the cells, which the arrowheads are pointing at are not nuclei. They are crowns of microvilli surrounding the cilium, - a similar picture is shown in Fig. 1 of the Momose et al., 2012 on Clytia, and on your own Fig 4L in the 48 hpa image. Nuclei are much bigger - see DAPI.\nWe thank the reviewer for pointing this out and for clarification, we have revisited our images and accordingly replaced the images in Figure S8D. We also added the missing “D” in this Figure.\nAlso, the authors show significant depletion of the mCherry signal in the cell contacts in iCRT14 in comparison with DMSO, which should not happen, since iCRT14 prevents b-cat/Tcf interaction, not something else.\nIndeed, the mCherry staining in the membrane seems also affected, however, we would not claim it to be significant as we did not perform any quantification. One explanation of this observation might be connected to the RNAseq data we obtained following iCRT14 treatments that revealed that ECM associated genes were downregulated under those conditions (Fig. 7C). Thus, rather than a direct effect of iCRT14 on the membrane pool of ß-catenin, the reduced mCherry staining at the membrane after iCRT14 treatments might be indirect and associated with reduced ECM associated genes, involved in cell-cell interactions.\nAnother point: I am not sure what “a”, “b”, “abc”, “ac” and “d” mean on top of the error bars on (C), but to me this plot shows that AZ is the only chemical with a clear effect on b-cat signaling. iCRT14 does not have a significant effect on b-cat in comparison to DMSO. It is not clear to me, how iCRT14 can abrogate the effect of AZ if it does not have an effect on topflash-driven expression when administered alone. A good control for the functionality of iCRT14 would be qPCR or RNA-Seq on b-cat target genes during regeneration. It should be reciprocal in iCRT14 and AZ - at least for genes upregulated in AZ.\nThe letters (“a”, “b”, “abc”, “ac”, “d”) above the error bars on (C) indicate groups that are statistically different based on the post-hoc test, as detailed in Supplementary Table 26. We added this information also in the legend of the corresponding figure. “Different letters above the bars indicate statistically significant differences among treatments (p < 0.05) according to two-sided pairwise Kruskal–Wallis tests followed by post hoc Dunn tests with Benjamini–Hochberg adjustment. Statistical analyses are provided in Sup Table 26”\nWe don’t really grasp the rationale why iCRT14 can’t significantly abrogate the overactivation of ß-catenin (measured by luciferase activity) in AZ treated animals when treated together. This is what our data show.\nHowever, we agree that the observation that iCRT14 doesn’t visibly affect topflash activity under physiological conditions might be somewhat puzzling. We propose that the absence of significant reduction in this context might be explained by the faint/basal level of luciferase activity in DMSO treated controls that are not sufficient to detect any decrease using this technique under physiological condition. In line with this idea, AZ-treated embryos co-treated with iCRT14 reach the same levels of luciferase activity as DMSO or iCRT14 alone.\nAlthough previous studies (Marlow et al, 2013; Faltine-Gonzalez et al, 2023) that we cite in the manuscript have used iCRT14 in Nematostella to inhibit the cWnt pathway, we developed experimental procedures to visualize and show the effect of this pharmacological inhibitor on cWNT signaling either using Topflash-driven luciferase expression (Fig. S8C) and on endogenous ß-catenin localization directly (Fig.S8D) as well as on regeneration (Fig. 5,6,7). In summary:\n1 - iCRT14 prevents the effects of AZ induced luciferase expression\n2 - regeneration-induced nuclearization of the endogenous ß-catenin at the amputation site is prevented in iCRT14-treated polyps\n3 - The iCRT14 caused phenotype (i.e., block of the regeneration process at “stage 2”) is in line with the activation pattern of the endogenous ß-catenin starting between 12-20hpa.\nAll together, our complementary results are in line with an inhibitory effect of iCRT14 on endogenous ß-catenin / cWnt signaling. In our study, we have performed RNAseq on iCRT14 treated animals during regeneration (See Fig. 7). In our opinion, regarding the main question of our study, all our results do not warrant further RT-qPCR or RNAseq experiments on AZ treated animals during regeneration.\nLines 334-335: Clumsy sentence. MEK/ERK and cWnt do not block development and patterning. Experimental modulation of MEK/ERK and cWnt does.\nWe thank the reviewer for pointing this out. The sentenced has been changed to:\n\"Previous studies have shown that perturbation of MEK/ERK and cWnt perturbs embryonic development and patterning in N. vectensis.\"\nLines 352-353: The authors write “iCRT14 treated polyps did not display ß-catenin nuclearization at 24hpa (Fig.S8C)” – but their controls also do not show b-cat nuclearization - at least on Fig. S8. See comment to Line 333. There is clear nuclear b-cat-mCherry signal on Fig. 4L at 12 and 24hpa.\nThis comment has been addressed in a previous comment above. Following the reviewer’s advice we have re-assessed our images and replaced the ones from Fig. S8D that now shows the abrogation of endogenous ß-catenin nuclearisation following iCRT14 treatments, compared to DMSO control.\nLine 363: Replace “as” with “in comparison to”\nDone.\nLine 445: On Fig. 7C, the GO-terms are completely unreadable without a 5x zoom, and certainly not on the printout. If this is a main text figure, every part of it has to be readable on a standard page without zooming.\nWe have taken into account the reviewers comment and now provide a summary Figure for 7C (main figure) and have moved the GO terms to Figure S14 (supplementary figure) so that they are bigger and more readable.\nLines 451-461: How do the authors explain that MAPK signaling is upstream of Wnt, but the genes controlled by Wnt are not affected by MAPK inhibition?\nThe reviewer raises a good point that we have attempted to answer in our revised version (l.462-469). While the temporal aspect we stressed might be an explanation, we now propose two additional possible reasons that might be even more compelling: a) the presence of compensatory regulatory mechanisms that drive cWnt target genes during regeneration in N. vectensis independently of MEK/ERK signaling, or b) spatially distinct actions of MEK/ERK signaling that affect amputation-induced activation of cWNT signaling in one region (e.g., the amputation site BWE), while cWNT signaling activation can remain unaffected in another region (e.g., mesenteries).\nWe completed this paragraph in the latest revision with the above mentioned elements.\nLines 476-481: The authors write “Systematically comparing these pools of genes (emb_cWnt, emb_ERK, regen_spe) with the up- or down-regulated genes following inhibition of MEK/ERK (Tables S20,S21), apoptosis (Tables S22,S23), or cWnt (Tables S24,S25) during regeneration, we observed that MEK/ERK signaling during regeneration has the most widely extended control on gene expression for all three pools (Fig.8; Fig.S15).” This makes total sense, but doesn’t it contradict what they wrote on lines 451-461 (that target lists barely overlap)?\nThe two analyses mentioned by the reviewer are addressing different questions:\n(i) downstream targets of MEK/ERK, apoptosis or cWNT during regeneration (Figure 7), (ii) to what extent are genes that are either downstream targets of MEK/ERK or cWNT signaling during embryonic development, or that belong to the set of “regeneration-specific” genes, are controlled by either MEK/ERK, apoptosis or cWNT during regeneration (Figure 8). The results therefore provide complementary, not contradictory information.\nMore specifically, the statement about the downstream targets that barely overlap (l.451-461) makes reference to Fig. 7, i.e., the downstream targets whose expression are affected by U0126, Z-VAD or iCRT14 treatments during regeneration. Fig.8 compares the downstream targets identified in Fig.7 to (a) embryonic downstream targets of MEK/ERK signaling, (b) downstream targets of cWnt signaling and (c) the “regeneration-specific” genes that are up-regulated at 20hpa. These three pools of genes make reference to Fig. 3.\nThe latter comparison highlights that part of the embryonic MEK/ERK or embryonic cWnt downstream targets as well as some of the “regeneration-specific” targets are controlled by MEK/ERK signaling during regeneration and not by cWNT or apoptosis.\nTo clarify this point, we have added the reference to Fig. 7 and Fig. 3 to the manuscript.\nLines 495-502: I do not find labels like Bi, Bii, and Biii on Fig. 8. I also do not see any insets on Fig. 7B.\nThe reviewer is correct and as the panels of Fig. 8 contain specific subtitles, we have removed the indications “i,ii and iii” in the manuscript. The insets are in Fig. 8B and we fixed this in the manuscript.\nLine 514: This is not a GRN. Unlike anything published by Eric Davidson, who was using the same type of graphics for GRN representation, here, it is just a list of downstream targets of MEK/ERK without any experimentally proven or even hypothetical regulatory links between them. It occupies figure space without providing any information a supplementary table cannot provide. Please remove.\nWe agree with the reviewer that the GRN blueprint we established, shown in Fig. 8D, does not include any regulatory interactions between the MEK/ERK downstream targets. We use the term GRN blueprint (Röttinger et al, 2012; Amiel et al, 2017) to indicate putative transcriptional interactions and clearly state in the Figure legend that no assumption on direct or indirect targets are made.\nL.547-548 further states: “This GRN blueprint indicates the potential direct or indirect downstream targets of MEK/ERK signaling at the onset of regeneration”. We have changed the term “GRN” by “GRN blueprint” in this sentence. We decided to keep this diagram / GRN blueprint as it a) not only provides a clear visual on the MEK/ERK downstream target genes during regeneration but b) also includes spatial information of these downstream targets. This GRN blueprint diagram might be of usage to the community for developing and testing regulatory hypotheses in follow-up studies.\nLine 534-536: The authors write that their results “support a model in which regeneration reuses core developmental programs such as cell type specification and patterning, while also activating unique, regeneration-specific regulatory elements and transcription factors”. They may be right in principle, but their results do not show anything of this sort. Out of 123 “regeneration-specific genes” in Table S1, there is a single transcription factor, Msx2-like, however, single-cell transcriptomics atlas of Cole et al., 2024 shows that it is strongly expressed in the retractor muscle and also in one particular neuroglandular cell type in a normal adult polyp. So, it may be “regeneration-specific” only in the comparison made by Andreoni-Pham et al., but not otherwise. Since there is no ATAC-Seq data in the manuscript, the authors cannot say that their data supports the existence of “regeneration-specific regulatory elements” either.\nIt seems that the way we formulated the sentence cited by the reviewer might lead to confusion in regard to what we defined as “regeneration-specific” genes versus the “regeneration-specific regulatory elements and transcription factors”. Only the latter is relevant to the cited sentence above.\nTo clarify this point, the revised sentence (changes are in blue) now reads:\n“Globally, our results support a model in which regeneration reuses core developmental programs such as cell type specification, proliferation and patterning, and suggest that regeneration activates gene expression via regeneration-specific regulatory elements”.\nThe rationale for this sentence is the following. Fig. 1, Fig. 2 and associated Supplementary Figures, show that there is a large overlap of genes dynamically expressed between embryogenesis and regeneration, and that some of these genes have been associated with cell type specification, patterning & proliferation.\nIn addition, Fig. 2G,H highlights for example the expression dynamics of the transcription factors tcf and runx. During embryonic development, their temporal expression patterns are different and they belong to different embryonic expression clusters (EX and EX), suggesting that they are regulated by different elements.\nInterestingly, these two TFs are co-expressed during regeneration (i.e., they share a highly similar temporal expression profile and belong to the same regeneration expression cluster R6), suggesting that they are regulated by the same regeneration-specific elements.\nLine 657: wnt7\nDone\nLine 703: The scheme on Fig. 9 is incorrect. MEK/ERK is responsible for the initiation of the endomesodermal expression in the plate in the 6-8 hpf blastula stage (Haillot et al., 2025); that is the same time when maternal beta-catenin goes into the nuclei of all future ectodermal cells (Lebedeva et al., 2025). So, there has to be not only the orange but also the green arrows to each embryo cartoon.\nWe thank the reviewer for pointing this out. However, rather than incorrect, we agree that the scheme on Fig.9 was missing an element. Following the reference cited by the reviewer we have added a green arrow to the blastula stage (no indication of MEK/ERK activity at the cleavage stage is indicated in Halliot et al, 2025). We also added a green arrow to the post-metamorphic juveniles stage as MEK/ERK signaling is required for metamorphosis (Rentzsch et al, 2008)."}]

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