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Regeneration programs buffer genetic defects in animal development [1]

['Kazunori Ando', 'Morgridge Institute For Research', 'Madison', 'Wisconsin', 'United States Of America', 'Department Of Cell', 'Regenerative Biology', 'University Of Wisconsin--Madison', 'Sushant Bangru', 'Department Of Cell Biology']

Date: 2026-09 

Regeneration programs enable animals to restore damaged or lost tissues, and the range of stimuli for these programs is incompletely understood. Here, we used zebrafish, a vertebrate species with exceptional regenerative capacity, to identify chemically induced mutations that alter regeneration-associated gene activation. Transgenic zebrafish with a permissive promoter and EGFP cassette inserted in the vicinity of the pro-regenerative factor gene fgf20a were mutagenized, and larvae homozygous for ENU-induced mutations were assessed for disruptions in fgf20a-directed reporter gene expression following fin fold amputation. One line was identified with heritable, elevated fgf20a:EGFP presence in the absence of experimental injury, localized to regions of fin fold tissue undergoing degeneration. Whole-genome sequencing (WGS) identified a mutation within exon 72 of the fraser syndrome 1 (fras1) gene, mutated in patients with inherited skin disease. fras1 mutant larvae spontaneously displayed broader signatures of regeneration, and zebrafish crispants for homologs of other genes mutated in human developmental diseases also displayed regeneration-associated gene expression in regions of dysmorphology. Tempering Fgf signaling by transgenic expression of a dominant-negative Fgf receptor in fras1 mutants exacerbated the disease phenotype. Our findings provide evidence that regeneration programs are harnessed in response to developmental defects caused by genetic mutations, potentially buffering deleterious phenotypes.

Funding: This work was supported by NIH Grant R01 HD105033 (to K.D.P.); awards from MEXT, Japan (15K21751) and the Uehara Memorial Foundation (to K.A.); and a Duke CAGT genome technology postdoctoral fellowship (to S.B.). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Copyright: © 2026 Ando et al. This is an open access article distributed under the terms of the Creative Commons Attribution License , which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Here, we conducted a forward genetic screen using a zebrafish fgf20a enhancer trap line to find mutations that disrupt activation of regeneration-associated gene expression upon amputation injury in larval fins. Unexpectedly, we identified a line with recessive mutations in fras1, a gene previously linked to epithelial integrity and extracellular matrix (ECM) organization in species from zebrafish to humans, causing spontaneous activation of fgf20a regulatory sequences. To explore a hypothesis that enhancer-driven regenerative networks are deployed in response to intrinsic developmental defects, we analyzed additional regeneration-responsive gene expression in these mutants, performed crispant analysis of other genes associated with human developmental disorders, and experimentally limited Fgf signaling during fras1 deficiency. Our results suggest a new and potentially important role for injury-induced gene expression programs as protective systems to buffer the effects of deleterious gene mutations.

Fluctuations in signals with impact during regeneration also occur during normal development and tissue homeostasis. Tissue growth, turnover, and morphogenesis naturally involve episodes of hypoxia, mechanical strain, inflammation, and epithelial remodeling. The zebrafish heart, for example, undergoes a period of strain during the juvenile stage that initiates a program with several commonalities as injury-induced heart regeneration in adults [ 26 ]. During lung branching morphogenesis, rhythmic tissue expansion generates epithelial tension that activates YAP signaling to drive proliferation and epithelial remodeling—mechanisms like those redeployed during adult lung repair and regeneration [ 27 ]. Likewise, physiological hypoxia serves as a morphogenetic cue during early embryogenesis, where low oxygen tension enhances WNT pathway activity and primitive-streak gene expression through HIF-dependent regulation [ 28 ]. Thus, regeneration programs have the potential to act as surveillance systems, engaged at any point in life when tissues experience stress or compromised integrity—even during early development.

A wide range of injury-associated cues can trigger regeneration programs. Reactive oxygen species, particularly hydrogen peroxide, form tissue-scale gradients within minutes of wounding and recruit leukocytes to initiate regenerative response [ 15 ]. Innate immune activity is similarly critical: macrophage-derived cytokines regulate blastema formation and patterning in both salamander limbs and zebrafish fins [ 16 , 17 ]. Oxygen levels provide another axis of control, as hypoxia can influence cardiomyocyte proliferation in zebrafish and neonatal mice [ 18 , 19 ]. Hormones are thought to gate regenerative competence; presence and activity of thyroid hormone levels suppress the capacity for cardiac regeneration, whereas disruption of thyroid hormone receptor signaling can enhance repair by altering metabolic and hypoxia pathways [ 20 – 22 ]. Innervation and mechanical strain are also early influences on regeneration [ 23 – 25 ]. Each of these triggering signals ultimately converges on regulatory controls like enhancers.

One of the most striking biological disparities between vertebrate species is their varied capacity to regenerate damaged tissues. Whereas many adult mammalian tissues display fibrosis and incomplete repair in response to injury, organisms like salamanders and zebrafish can regenerate fins, limbs, heart, spinal cord, kidney, and other tissues/organs with remarkable fidelity [ 1 , 2 ]. Across these diverse tissue contexts, regeneration relies on establishing transcriptional signatures that trigger cell proliferation, direct tissue architecture, and restore tissue function. Broadly, these changes in gene expression that occur during regeneration, versus what normally is in place in the absence of injury, are referred to as “regeneration programs”. In recent years, it has become clear that regeneration programs are orchestrated by dedicated cis-regulatory elements [ 3 , 4 ]. First referred to as tissue regeneration enhancer elements (TREEs) and also called damage-responsive elements or regeneration-responsive elements, these sequences receive injury signals and selectively activate transcription in damaged tissue [ 3 , 5 , 6 ]. For example, the Fgf ligand gene fgf20a is induced within hours of fin amputation in zebrafish, where it is required to initiate formation of the regeneration blastema [ 7 , 8 ], and multiple TREEs appear to collectively direct its contextual expression [ 9 ]. Enhancers near leptin, il11, and inhba genes similarly direct target gene transcription after fish fin amputation [ 3 , 6 , 10 ]. Many other TREEs have been identified across diverse species, tissues, cells, and injury contexts [ 11 – 14 ].

Results

ENU mutagenesis screen for mutations that alter regeneration-associated fgf20a expression Enhancer trap and reporter lines for injury-responsive genes enable real-time visualization of regeneration-associated gene expression and can be powerful tools for dissecting the molecular logic of regeneration (Fig 1A). To identify potential regulatory factors required for an essential factor for fin regeneration, fgf20a, we performed ENU mutagenesis in homozygous males of an fgf20a enhancer trap line (HGn21A, or fgf20a:EGFP; Fig 1B) [29]. In this line, a transposon-based hsp70l:EGFP cassette randomly inserted 1.5 kb upstream of the fgf20a gene locus. fgf20a:EGFP animals display strong, injury-induced EGFP expression after caudal fin amputation or larval fin fold amputation, mimicking endogenous fgf20a expression [30]. We treated 11 homozygous fgf20a:EGFP male fish (F 0 ) with ENU, and, after confirming a high mutagenesis rate, we prepared 117 F 2 families, all homozygous for fgf20a:EGFP, to breeding age by intercrossing 190 F 1 fish (Fig 1C). Larval fin fold regeneration is robust in zebrafish, and both fgf20a mRNA and fgf20a:EGFP are induced within one day of amputation of the 3 day post fertilization (dpf) larval fin fold (Fig 1D) [8]. In total, we screened 9,868 total animals from 460 F3 families (all homozygous for fgf20a:EGFP and for ENU-induced mutations) for induction of fgf20a:EGFP after larval fin fold amputation. PNG larger image

TIFF original image Download: Fig 1. ENU mutagenesis identifies a mutation that increases fgf20a-directed expression at the degenerating fin fold. (A) Regeneration-responsive genes are regulated by tissue regeneration enhancer elements (TREEs) and tissue regeneration silencer elements (TRSEs). (B) Regeneration-responsive expression of the pro-regenerative factor, fgf20a, is regulated by a TREE, fgf20aE48, as present in the enhancer trap transgenic reporter line, HGn21A. (C) ENU mutagenesis screen flow. Homozygous male HGn21A (fgf20a:EGFP) fish were treated with ENU and mated with homozygous fgf20a:EGFP females to generate F 1 families. F 2 families were generated from intercrossing F 1 fish, and screens of F 3 larvae were performed to detect homozygous mutations that reduce or increase injury-induced fluorescence (m 1 , m 2 , m 3 ,...). (D) (Top) Uninjured 4 dpf larva. (Bottom) 4 dpf larva after fin fold amputation at 3 dpf. EGFP expression is clearly induced at the injury site (yellow arrowhead in the right magnified view of tail region), mimicking fgf20a induction after the same injury. (E) The heritable mutant larvae (budlight) identified from screens spontaneously displayed a shortened body (yellow double-headed arrow) and elevated expression of fgf20a:EGFP in the degenerated fin fold (yellow arrows). (F) Quantification of relative expression levels of fgf20a in wild-type uninjured larvae, wild-type larvae at 1 day post-amputation (dpa), and budlight mutants at 4 dpf. Expression was significantly increased in wild-type larvae following injury and was highest in budlight mutants. Statistical significance was determined using a two-sample t test assuming unequal variances. P values are indicated above brackets. Data represent mean ± SEM. n = 3 pooled biological replicates per condition, with 20 larval tails pooled per replicate. Scale bars, 100 μm. The data underlying the graphs can be found at https://datadryad.org/search?q=10.5061%2Fdryad.jdfn2z3sm. https://doi.org/10.1371/journal.pbio.3004011.g001 From these 460 families, 144 contained larvae with potential changes in fgf20a:EGFP reporter expression and/or defects in regeneration. In reassessments of these families, 11 displayed abnormal regeneration 1 day after fin fold amputation. Many families contained larvae displaying limited fin fold regeneration, but also with developmental defects like fin fold degeneration, cardiac edema, craniofacial malformation, or unusual trunk curvature (S1A and S1B Fig). Two of these families had members displaying subtle defects in fin regeneration at the adult stage -- apparent delays in initiating regeneration but recovery to normal regeneration (S1C Fig). Interestingly, while several families had increased fgf20a:EGFP in fin fold or trunk muscle, all of these families showed roughly normal fgf20a:EGFP expression at the amputation site. Only one of the families had members displaying a gross change in fin fold fgf20a:EGFP from clutchmates. These larvae displayed a jagged, degenerated fin fold in the absence of injury along with a conspicuous increase in fgf20a:EGFP at the caudal and ventral fin folds, and in pectoral fin buds, and thus we referred initially to this mutant as budlight (Fig 1E). qPCR analysis revealed that endogenous fgf20a levels are many-fold higher in budlight mutant tails than those of clutchmates (Fig 1F). To quantitatively assess fin fold degeneration and regeneration-associated signaling in budlight mutants, we measured fin fold area and EGFP reporter activity across larval development from 1–6 dpf (S2A and S2B Fig). Wild-type larvae displayed progressive expansion of fin fold tissue during normal development, whereas budlight mutants exhibited a pronounced failure of fin fold growth beginning around 2–3 dpf, resulting in reduced tissue area at later stages (S2A Fig). Concomitant with this degeneration phenotype, EGFP reporter intensity, normalized to tissue area, increased sharply in budlight mutants but remained low in wild-type larvae (S2B Fig). Notably, reporter activation coincided with the onset of overt tissue degeneration, suggesting that injury/regeneration-associated signaling programs are activated early during mutant fin fold degeneration. In summary, ENU mutagenesis revealed a heritable mutation that causes spontaneous elevation of the fgf20a regeneration program in uninjured fin primordia undergoing degeneration.

A nonsense mutation in fras1 causes ectopic fgf20a-directed gene expression To identify the molecular basis of budlight, we performed whole-genome sequencing using genomic DNA from 5 dpf wild-type larvae homozygous for the fgf20a:EGFP insertion, as well as from budlight siblings displaying the degenerating fin fold phenotype (Fig 2A). Sequence reads were aligned to the zebrafish reference genome, and we applied filtering steps to remove background variants, sequencing artifacts, and low-quality calls. This filtering enriched for homozygous variants unique to the mutant pool, thereby prioritizing candidate lesions most likely to underlie the phenotype. SNP homozygosity analysis revealed a single point mutation in exon 72 of the fraser extracellular matrix complex subunit 1 (fras1) gene, corresponding to the third-to-last exon. This variant represented a nonsense mutation (T to A), introducing a premature stop codon in place of tyrosine at amino acid position 3,708 (Fig 2B). PNG larger image

TIFF original image Download: Fig 2. Identification of a nonsense mutation in fras1, and similar effects on fgf20a expression following genetic disruption of developmental disease-associated genes. (A) Workflow schematic for whole-genome sequencing-based identification of the causal mutation underlying the budlight phenotype. Homozygous mutant larvae displaying elevated fgf20a:EGFP expression were isolated from budlight heterozygous crosses at 5 dpf and pooled for genomic DNA extraction, library preparation, and Illumina sequencing. Sequence reads were aligned to the zebrafish reference genome followed by variant calling and identification of homozygous candidate mutations. (B) Whole-genome sequencing identified a nonsense mutation in fras1 consisting of a T-to-A substitution in exon 72, converting a tyrosine codon into a premature stop codon. Representative sequencing alignments from mutant and sibling control samples are shown. (C) CRISPR-Cas9 F 0 mutagenesis targeting fras1 phenocopied the budlight mutant phenotype and induced ectopic fgf20a:EGFP expression. Representative examples of weak and strong reporter activation are shown in degenerating fin fold and craniofacial tissues. Control embryos displayed only canonical developmental fgf20a:EGFP expression domains. (D–G) Representative examples of phenotypes and ectopic fgf20a:EGFP activation observed following CRISPR-Cas9 F 0 mutagenesis of zebrafish orthologues of human developmental disease-associated genes, including pex1 (D), tbx5a (E), grip1a (F), and frem2a (G). Yellow arrows indicate regions with weak fgf20a:EGFP activation, and green arrows indicate regions with strong reporter induction. (H) Quantification of phenotypic classes observed in the CRISPR-Cas9 F 0 screen. Larvae were categorized as displaying no detectable phenotype, minor defects with ectopic fgf20a:EGFP expression, or major defects with ectopic fgf20a:EGFP expression. Statistical significance was assessed using Fisher’s exact test with multiple testing correction. *p < 0.05, **p < 0.01. The data underlying the graphs can be found at https://datadryad.org/search?q=10.5061%2Fdryad.jdfn2z3sm. https://doi.org/10.1371/journal.pbio.3004011.g002 fras1 encodes a transmembrane protein with domains including von Willebrand factor–C and furin-like repeats, and it is crucial for epithelial–mesenchymal adhesion and basement membrane stability. In humans, mutations in FRAS1 cause Fraser syndrome, an autosomal recessive developmental disorder marked by epidermal blistering, syndactyly, and renal anomalies [31]. In zebrafish, fras1 is expressed in epithelial tissues, particularly in developing fin folds, and is required for skin integrity and craniofacial morphogenesis [32]. Additionally, zebrafish mutants in fras1, as well as orthologues of FREM1/2 and other components of the Fraser syndrome complex, display classic fin blistering phenotypes that recapitulate aspects of the human and mouse diseases [33]. We observed blistering in fras1 mutant fin fold at 1–2 dpf and not thereafter (S2C Fig), consistent with the defects of Fraser syndrome. We do not resolve whether posterior fgf20a:EGFP expression, and other responses described later, are responses to blister resolution and/or progressive degeneration of the affected tissue. To independently test whether fras1 disruption was responsible for the fras1pd416 degenerative phenotype and activation of the fgf20a:EGFP enhancer trap, we designed CRISPR/Cas9 guides targeting fras1 coding sequences. Injection of gRNAs into one-cell stage embryos generated mosaic F 0 animals that phenocopied the ENU mutant: at 3 dpf, injected embryos exhibited localized fin fold degeneration accompanied by ectopic fgf20a induction in the pectoral fin bud, ventral fin fold, and caudal fin fold (Fig 2C). These CRISPR experiments establish a causal link between fras1 loss-of-function and the spontaneous activation of regeneration programs observed in budlight mutants, which we refer to hereafter as fras1 mutants.

Crispants in zebrafish orthologues of human congenital disorder genes show ectopic fgf20a-directed gene expression Based on the association of fras1 mutations with fgf20a induction, we postulated that, more broadly, intrinsic defects in tissue architecture that arise from genetic lesions in developmentally key genes can trigger regeneration programs. For example, intrinsic fragility or biomechanical stress as the result of mutations could activate regeneration programs that act as surveillance mechanisms. One prediction of this idea is that other genetic defects beyond fras1 mutations should cause induction of regeneration-associated fgf20a expression. To test if activation of regeneration-associated transcription is a general response to developmental defects, we performed a targeted loss-of-function (LOF) screen in the fgf20a:EGFP enhancer trap background. We selected 15 zebrafish orthologues of human disease genes that, when mutated in humans, cause congenital syndromes involving the skin, skeleton, or craniofacial structures—tissues related to those where fgf20a is induced by experimental injury (S3A Fig). These included genes linked to Fraser syndromes (fras1, frem2, grip1), osteogenesis imperfecta and related bone fragility (bmp1a, col1a1, col1a2, col2a1), skeletal dysplasia (fgfr3, flnb, nek1, ebp), limb and heart malformations (tbx5a), and cutaneous or connective tissue disorders (st14, lmna, pex1, aspa). We chose these genes based on documented human pathologies and evidence from zebrafish models: for instance, fras1/pinfin and frem2/blasen mutants present fin blistering [33], bmp1a mutants have ‘frilly fins’ owing to deficient collagen processing [34], tbx5a/heartstrings mutants exhibit pectoral fin and heart malformations [35], zebrafish fgfr3 mutants show craniofacial malformations and delayed ossification similar to human skeletal dysplasia [36], and CRISPR-created imna mutants present muscle and mobility defects analogous to muscular laminopathies [37]. Injection of CRISPR/Cas9 reagents targeting these genes resulted in larvae with varying degrees of morphological defects. In nearly every case where fin fold tissue integrity was visibly compromised, we observed ectopic activation of the fgf20a:EGFP enhancer trap in the affected regions (Figs 2D–2G, S3C and S3D). The induced EGFP expression domains extended beyond the normal developmental patterns of fgf20a, which typically include trunk muscle and craniofacial structures, and instead appeared in affected fin fold epithelia. Larvae that had been injected as embryos with Cas9 protein with or without non-targeting guide RNAs did not exhibit ectopic fgf20a:EGFP activation or overt tissue degeneration, indicating causal effects of specific gRNAs (S3B–S3D Fig). Statistical analyses indicated that approximately half of the tested genes produced a significant enrichment of larvae in the “defect plus ectopic fgf20a expression” category (Fig 2H). To complement the qualitative scoring of ectopic fgf20a:EGFP activation in F0 CRISPR embryos, we quantified fin fold tissue area and reporter activity at 4 dpf. Area measurements normalized to control larvae revealed that several perturbations identified by qualitative assessment as exhibiting strong enhancer activation and morphological defects also displayed significant tissue loss (S3E Fig). In parallel, integrated EGFP intensity normalized to fin fold area was elevated across most perturbations associated with ectopic reporter activation (S3F Fig), indicating enrichment of fgf20a:EGFP signal within the remaining fin fold tissue. Together, these quantitative analyses support the qualitative observation that developmental perturbations associated with tissue degeneration are accompanied by activation of fgf20a expression.

Ectopic activation of broad regeneration responses in fras1 mutant larvae Another prediction of our hypothesis is that loss of fras1 function should trigger regeneration-responsive transcriptional responses in addition to fgf20a. To test this prediction, we focused on two well-characterized TREEs, the first of which is LEN, which is required for regeneration-associated expression of the leptinb gene and directs expression of reporter transgenes in regenerating fin folds, adult fins, and adult hearts [3]. The second is REN, which is linked to the runx1 locus and directs expression of reporter transgenes in injured and regenerating hearts; we found REN also directs regeneration-responsive expression in mesenchyme and notochord of RENcfos:EGFP larvae (Fig 3A and 3B) [3,38]. To test whether fras1-associated degeneration is sufficient to engage TREEs, we crossed LENP2:EGFP or RENcfos:EGFP reporter transgenes into the fras1 background. We found that fras1 mutants exhibited robust, ectopic activation of both reporters in affected fin fold tissue by 4 dpf, with patterns resembling their normal injury-induced expression domains (Fig 3C and 3D). This response was observed in all mutant larvae examined, whereas control siblings showed no detectable reporter expression in intact fin folds. PNG larger image

TIFF original image Download: Fig 3. Evidence for a broad regeneration program activated in fras1 mutants. (A) TREE reporter lines with runx1 linked enhancer (REN) and leptin b linked enhancer (LEN) upstream of minimal promoter-EGFP cassettes. cfos, a 100 bp promoter of mouse cfos. P2, a 2 kb promoter of zebrafish leptin b. (B) Expression of reporters, fgf20a:EGFP, RENcfos:EGFP and LENP2:EGFP in regenerating larval fins at 4 dpf, after fin amputation at 3 dpf. fgf20a:EGFP has regeneration-responsive expression in epithelia, mesenchyme, notochord, and muscle, while RENcfos:EGFP and LENP2:EGFP, have regeneration-responsive expression in notochord and mesenchyme. (C) fras1 mutations induce EGFP expression in notochord and mesenchyme (yellow arrows) of RENcfos:EGFP larvae and in mesenchyme (yellow arrows) of LENP2:EGFP larvae. (D) Measurement of EGFP-positive area at distal regions of 4 dpf tails in uninjured wild type (white bars), 1 dpa wild type (gray bars) and uninjured fras1 mutants (red bars) with RENcfos:EGFP or LENP2:EGFP transgenes. Individual dots indicate biological replicates (n = 3–7). Data are presented as mean ± SEM. Statistical significance was assessed using two-way ANOVA. P values are shown above the corresponding comparisons. (E) Principal component analysis (PCA) of bulk RNA-seq datasets generated from wild-type (WT) uninjured larvae (4 dpf), wild-type regenerating tails at 1 and 2 dpa, and age-matched fras1 mutant larvae. PC1 and PC2 account for 40.1% and 33.6% of total variance, respectively. (F) Sample coordinates across the first four principal components. Percentage variance explained for each principal component is indicated below the corresponding axis. (G) Correlation analysis comparing transcriptional changes during wild-type regeneration (1 dpa vs. wild-type uninjured) and fras1 mutants (fras1 vs. wild-type uninjured). Each point represents a single gene with |log 2 FC| > 1 during wild-type regeneration. Genes changing concordantly between regeneration and fras1 mutants are shown in orange, while discordantly regulated genes are shown in blue. Representative regeneration-associated genes are labeled. Pearson and Spearman correlation coefficients are indicated. Shaded diagonal region denotes concordant fold-change directionality. (H) Heatmap showing transcriptional dynamics of representative regeneration-associated and fras1 mutant-specific gene sets across wild-type uninjured, wild-type 1 dpa, wild-type 2 dpa, and fras1 mutant conditions. Gene expression values are row-scaled z-scores. Adjacent panels summarize enriched Gene Ontology categories associated with each gene set. Displayed categories represent representative nonredundant functional groups selected after collapsing highly overlapping and related GO Biological Process terms. Complete gene-level differential-expression and GO-term assignments are provided in S3 Table. The data underlying the graphs can be found at https://datadryad.org/search?q=10.5061%2Fdryad.jdfn2z3sm. https://doi.org/10.1371/journal.pbio.3004011.g003 To broadly test whether the fin fold degeneration observed in fras1 mutants is associated with activation of regeneration programs, we performed bulk RNA-seq on isolated tail tissues from uninjured larvae (4 dpf), wild-type regenerating tails at 1 and 2 days post-amputation (dpa), and age-matched fras1 mutant larvae at 4 dpf (S4A Fig). Principal component analysis (PCA) revealed that fras1 transcriptomes clustered separately from uninjured wild-type controls and partially aligned with regenerating wild-type tissues along PC1, which accounted for 40.1% of total variance (Fig 3E and 3F). In contrast, PC2 (33.6% variance explained) distinguished regenerating wild-type samples from fras1 mutants, suggesting that while the mutant engages regeneration-associated transcriptional programs, it also exhibits distinct transcriptional features. Consistent with this interpretation, gene ontology (GO) analysis of genes contributing to PC1 identified enrichment for regeneration-associated processes including ECM organization, connective tissue development, regeneration, and appendage morphogenesis, whereas PC2-associated genes were enriched for developmental and sensory perception pathways (S4B Fig). To directly assess the relationship between regeneration and mutant transcriptional states, we compared gene expression changes between wild-type regeneration (1 dpa versus uninjured wild-type) and fras1 mutants (fras1 versus uninjured wild-type). Genes strongly altered during early regeneration exhibited a high degree of concordant regulation in fras1 mutants (Pearson r = 0.8; Spearman ρ = 0.8; Fig 3G). Notably, 90% of regeneration-associated genes changed in the same direction in fras1 mutants (Q1 + Q3 quadrants), including known regeneration-associated genes such as lepb, fgf20a, il1b, tnfb, and shhb. These findings suggest that fras1 mutants spontaneously activate substantial components of the transcriptional program for regeneration in the absence of physical injury. To further resolve shared and mutant-specific transcriptional responses, we classified genes based on whether they were similarly regulated during wild-type regeneration and in fras1 mutants. Approximately 52.5% of transcriptionally altered genes were shared between regeneration and fras1 conditions, while 25.9% represented fras1-specific changes and 21.6% were regeneration-associated genes with limited activation in the mutant background (S4C Fig). Heatmap analysis indicated that regeneration-associated genes were broadly upregulated in fras1 mutants, particularly inflammatory and cytokine-associated pathways, whereas distinct fras1-specific programs included ECM organization, skin development, and metabolic pathways (Fig 3H). GO analysis further revealed enrichment of inflammatory response, cytokine-mediated signaling, and acute-phase response pathways among both regeneration-induced and fras1-downregulated gene sets, consistent with persistent activation of injury-associated signaling networks in the mutant state (Fig 3H). Together, these findings argue that loss of fras1 gene function induces a transcriptional state that partially recapitulates injury-induced regeneration programs while also engaging mutant-specific developmental and ECM remodeling pathways.

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