A macrophage-induced subpopulation of mesenchymal cells expressing Fcer1g contributes to wound-induced fibrosis

Domain: neuroscience
Published: 2026-02-12
Editorial outcome: minor_revision

Abstract
Fibrosis commonly occurs during adult skin wound healing, characterized by excessive extracellular matrix (ECM), leading to scarring. Mesenchymal cells, the primary ECM-producing population, are heterogeneous with varying fibrotic propensity during healing. While pro-fibrotic embryonically derived mesenchymal lineages have been identified, adult mesenchymal cells responsible for fibrosis are not yet fully characterized. In adult mice with conditional macrophage depletion during the early phase of wound healing, wounds exhibit attenuated fibrosis and a reduction in mesenchymal cell numbers. Here we show that early phase macrophage induces a distinct PDGFRα⁺ mesenchymal population expressing Fcer1g. This cell population expands rapidly after injury, shows high proliferative activity, and is largely absent when macrophages are depleted. Targeted ablation of this cell population does not delay wound closure but results in diminished scarring. Human wound datasets identified a transcriptionally conserved FCER1G-expressing mesenchymal subset, suggesting that this pro-fibrotic mesenchymal state is preserved in human wound healing.

Introduction
Following full-thickness injury, most adult mammalian skin undergoes a repair process that is accompanied by fibrosis. Wound healing is a highly coordinated, multicellular process progressing through sequential stages: inflammation, proliferation and remodeling1. The early phase of wound repair (day 0–5) is characterized by inflammation, during which the immune response is activated2. Immune cells, including macrophages, are rapidly recruited to the injury site, where they clear debris, combat infection, and initiate signaling cascades for tissue repair. Besides this, increasing evidence supports the correlation between macrophages and fibrotic responses: early fetal wounds, which recruit minimal macrophages, heal with negligible fibrosis3; similarly, wounds in neonatal PU.1-null mice, which lack macrophages and neutrophils, heal without fibrosis4; in adult LysMCre;iDTR mice, macrophage depletion during the early phase of wound healing leads to delayed wound closure but significantly reduces fibrosis5. These findings suggest that macrophages play an important role in modulating fibrosis.

The proliferative phase of wound healing (day 3-14) is marked by the formation of granulation tissue, characterized by aggregation of mesenchymal cells and deposition of ECM components6–8. This provisional ECM undergoes remodeling and contributes to scar formation. During the healing process, mesenchymal cells are the main driver of fibrosis as they are the predominant cell type synthesizes and remodels the ECM9. Prior lineage tracing experiments revealed the heterogeneity among mesenchymal cells10,11. Embryonic mesenchymal precursors expressing delta-like non-canonical Notch ligand 1 (Dlk1)12, Engrailed-1 (En1)13 or paired related homeobox 1 (Prrx1)14 generate pro-fibrotic mesenchymal lineages. Targeted ablation of these embryonic-derived lineages reduces cutaneous fibrosis without delaying wound closure13,14. These results support the concept that different mesenchymal subpopulations are independently responsible for pro-fibrotic or pro-regenerative roles.

One mechanism by which macrophages modulate fibrosis is through the regulation of mesenchymal cells. During wound healing, macrophages promote migration and proliferation of mesenchymal cells15. Early-phase macrophage depletion in LysMCre;iDTR reduces mesenchymal cell numbers and their proliferation5,15. In addition, macrophages activate the expression of ECM components in mesenchymal cells through signaling pathways such as TGF-β16 and interleukin-4 receptor α (IL-4Rα)17. However, the pro-fibrotic mesenchymal subpopulation regulated by macrophages remains to be explored.

In this work, we perform single-cell RNA sequencing (scRNA-seq) in Macrophage Fas-Induced Apoptosis (MaFIA) mice, in which macrophages are conditionally depleted during the early phase of wound healing. We identify a macrophage-induced mesenchymal subpopulation that expresses Fcer1g, expands early after injury, and contributes to fibrosis. We further show that targeted ablation of this population attenuates scar formation, identifying it as a key cellular driver of wound fibrosis.

Results
A subpopulation of mesenchymal cells is induced by early phase macrophages during wound healing
To conditionally deplete macrophages, we used the MaFIA mouse, which expresses a mutant human FAS death receptor and green fluorescent protein (GFP) under the control of the colony-stimulating factor-1 receptor (Csf1r) promoter18. AP20187 is a dimerizer that specifically activates the FAS death receptor to initiate cellular apoptosis. In vivo macrophage depletion during early wound healing (day 0–5) was achieved through daily AP20187 injections for five consecutive days, while control mice received vehicle treatment. On the final day of injection, we generated 5-mm full-thickness wounds on the dorsal skin of adult MaFIA mice using an excisional wound splinting model19,20. Macrophage depletion efficiency was evaluated three days later to reflect depletion status during the early healing response. GFP expression driven by the Csf1r promoter enabled flow cytometry–based quantification of macrophage depletion by measuring the percentage of GFP⁺ cells (Fig. 1a). At 3 days post injury (dpi), AP20187 treatment depleted at least 80% of GFP+ macrophages in the skin (Fig. 1b). To further confirm that GFP⁺ depletion reflects loss of macrophages, we performed flow cytometry for F4/80 and CD11b double-positive cells in the bone marrow, spleen, and skin from vehicle- and AP20187-treated MaFIA mice (Supplementary Fig. 1). These analyses confirmed robust depletion of F4/80⁺ CD11b⁺ macrophages in the skin (>75%) (Supplementary Fig. 1i, j) and moderate depletion in the bone marrow and spleen (~50%) (Supplementary Fig. 1a, b, e, f). Immunofluorescence staining further validated the loss of F4/80⁺ macrophages in these tissues (Supplementary Fig. 1c, d, g, h, k, l). By 7 dpi, F4/80+ macrophages began to repopulate in the skin18 (Supplementary Fig. 1m–p).

We then examined the wound healing outcomes. Consistent with the previous findings in adult LysMCre;iDTR mice5, early-phase macrophage depletion led to significantly reduced fibrosis at 21 dpi compared to vehicle-treated controls (Fig. 1c, d and Supplementary Fig. 2a). Masson’s trichrome staining revealed that AP20187-treated wounds exhibited smaller areas of thick, densely packed collagen fibers, indicating attenuated fibrosis (Fig. 1e and Supplementary Fig. 2b). Serial sections demonstrated reduced fibrosis in AP20187-treated wounds spanning from the wound center to the edge, as fibrosis was nearly absent at 150 μm from the center in the depleted group, while still clearly visible at 250 μm in controls (Supplementary Fig. 2c). Moreover, immunofluorescence staining for keratin 10 (K10) and CD31 revealed no significant differences in reepithelialization or angiogenesis between vehicle- and AP20187-treated wounds at 7 dpi and 14 dpi (Supplementary Fig. 3), indicating that the reduction in fibrotic area specifically reflects macrophage-dependent regulation of wound fibrosis. Previous studies have shown that hair follicles in the anagen phase can actively contribute to mouse wound re-epithelialization21, making it critical to ensure consistent hair cycle phases across experimental groups. The anagen phase is characterized by active cell proliferation22, as indicated by robust Ki67 expression (Supplementary Fig. 4b), and by activated Wnt/β-catenin signaling23, evidenced by nuclear β-catenin localization (Supplementary Fig. 5a). Around 7 weeks of age, follicles transition into the telogen phase, marked by the absence of both Ki67 and nuclear β-catenin, and remain in this phase until at least 12 weeks postnatal (Supplementary Figs. 4a, b and 5a). In our study, mice were wounded at approximately 8 weeks of age and analyzed around 11 weeks, during which time hair follicles are expected to be in the telogen phase. Consistent with this, both vehicle- and AP20187-treated groups showed no follicular Ki67 expression throughout the experimental timeline (Supplementary Fig. 4c), indicating limited proliferative activity. Moreover, the absence of nuclear β-catenin in both groups (Supplementary Fig. 5b) further confirmed that all animals remained in the telogen phase during the study period.

To identify mesenchymal subpopulations regulated by macrophages during early wound healing (day 0–5), we performed scRNA-seq on wound tissues from MaFIA mice with or without macrophage depletion at 3 dpi. This time point coincides with substantial macrophage infiltration and active mesenchymal cell proliferation24. UMAP analysis of the scRNA-seq data identified diverse cell populations, which were annotated using published murine skin scRNA-seq data and consensus marker genes from bulk transcriptomic studies11,25,26 (Fig. 1f, Supplementary Fig. 6 and Supplementary Table 1). The UMAP analysis identified populations including neutrophils (clusters 2, 5 and 15) expressing Mmp8, Retnlg, Efdc21, Ly6g, macrophages (clusters 0 and 1) expressing Pf4, Adgre1, Itgam, and Dendritic/Langerhans cells (clusters 11 and 20) expressing Cd74, Cd80, Cd86. Dendritic/Langerhans cells are a subset of tissue-resident macrophages with antigen-presenting function27, which were hereafter included as a macrophage population.

AP20187 administration significantly reduced macrophage populations (clusters 0, 1, 11, and 20) at 3 dpi (Supplementary Fig. 7a, b). This depletion altered the proportions of several cell types in the wound, including neutrophils, mesenchymal cells and epithelial cells (Supplementary Fig. 7c). This result suggests that macrophages regulate the activation or maintenance of these populations during healing. Among the identified clusters, clusters 4, 9, 14, 24, and 25 were classified as mesenchymal cells, because they expressed canonical mesenchymal markers, including Col1a1, Col1a2, Pdgfrα, Col5a1, Loxl1, Lum, Fbln1 and Cd3412,28,29. To further examine the specific mesenchymal subpopulation regulated by macrophages, we re-anchored, re-integrated and re-clustered the mesenchymal cells separately. This approach identified 16 mesenchymal subpopulations (Fig. 1g). Among them, subcluster 4 had high expression of Dpp4 (CD26), a marker associated with adipocyte precursor cells (APs)15 (Supplementary Fig. 8a). Subcluster 6 had high expression of Acta2 (smooth muscle actin), a hallmark of myofibroblasts30 (Supplementary Fig. 8b). The proportions of most mesenchymal subpopulations were not altered by macrophage depletion (Supplementary Fig. 8c). However, subcluster 9 was almost completely absent in wounds from macrophage-depleted mice (Fig. 1h and Supplementary Fig. 8c), suggesting a macrophage-dependent regulation of this mesenchymal subpopulation.

The subpopulation of mesenchymal cells induced by macrophages expresses Fcer1g
To identify unique gene markers that are both highly and broadly expressed in mesenchymal subcluster 9, we used the FindMarkers function in Seurat31,32. Comparisons were restricted to subcluster 9 versus other mesenchymal subclusters. Genes were evaluated based on expression level (log2 fold change) and prevalence (percentage of cells expressing the gene). Specifically, we calculated a uniqueness score (U-score) using the formula: U-score=log2 fold change × (percent expression in subcluster 9 − percent expression in all other subclusters), with criteria of log2 fold change ≥ 0 and adjusted p-value < 0.05. Using this approach, we identified several genes uniquely enriched in subcluster 9, including Fcer1g, Cd53, Lyz2, and Lcp1, with Fcer1g exhibiting the highest U-score (Fig. 2a, b and Supplementary Data File 1).

Fcer1g encodes the Fc fragment of the IgE receptor (FCER1G). scRNA-seq analysis revealed that among all mesenchymal subpopulations, Fcer1g-expressing mesenchymal cells uniquely expressed hematopoietic markers, particularly the myeloid-related marker Lyz2 (Fig. 2a). Previous studies reported that some wound-associated mesenchymal cells express myeloid-related genes11. Further analysis of published time-course scRNA-seq data on wound healing in mice confirmed that Fcer1g-expressing mesenchymal cells form a distinct cluster during early wound healing, co-expressing Tyrobp, Lyz2, and Lcp1 (Supplementary Fig. 9a)26.

To exclude the possibility that the reduction of Fcer1g-expressing mesenchymal cells was due to Csf1r-driven apoptosis, we examined the expression of Csf1r-driven Fas and GFP in this population. Since FCER1G is widely expressed in non-mesenchymal cells, such as T cells and macrophages33,34, and immunohistochemistry confirmed its expression in multiple immune cell types within the wound bed (Supplementary Fig. 9b), we used platelet-derived growth factor receptor alpha (PDGFRα) as an additional marker because PDGFRα is broadly expressed in dermal mesenchymal cells during wound healing35,36. Immunostaining of FCER1G+ PDGFRα+ mesenchymal cells in MaFIA wounds did not show GFP expression (Fig. 2c), indicating no Csf1r-driven mutant FAS receptor expression. Therefore, their reduction was not a direct consequence of AP20187-induced apoptosis but rather a result of interactions with macrophages.

We next used immunostaining to examine the spatial distribution of Fcer1g-expressing mesenchymal cells in unwounded and wounded skin, with wound bed and periwound strictly defined (Supplementary Fig. 10a). In unwounded skin, FCER1G+ PDGFRα+ mesenchymal cells accounted for less than 1% of the total cell population and less than 20% of total PDGFRα+ mesenchymal cells (Fig. 2d, e and Supplementary Fig. 10b, c). Following injury, their numbers increased significantly, first appearing in the periwound dermis at 3 dpi, peaking at 7 dpi, and gradually declining thereafter, plateauing at a level around two times of the unwounded skin (Fig. 2d and Supplementary Fig. 10b, c). By 7 dpi, this population was enriched in the wound bed with approximately 80% of PDGFRα+ cells expressing FCER1G (Fig. 2e and Supplementary Fig. 10b, c). From 7 dpi onward, FCER1G+ PDGFRα+ cells remained predominantly localized in the wound bed, with fewer cells in the periwound area (Fig. 2d and Supplementary Fig. 10c).

To assess the spatial relationship between FCER1G⁺ PDGFRα⁺ mesenchymal cells and vascular structures, we performed triple immunofluorescence staining for CD31, PDGFRα, and FCER1G. FCER1G⁺ PDGFRα⁺ cells were frequently located in proximity to CD31⁺ endothelial structures but did not colocalize with them, indicating that these mesenchymal cells are perivascularly associated rather than endothelial in identity (Supplementary Fig. 10d).

Having established the spatiotemporal dynamics of Fcer1g-expressing mesenchymal cells during wound healing, we next examined how their presence was affected by macrophage depletion. In AP20187-treated MaFIA mice, FCER1G+ PDGFRα+ mesenchymal cells were significantly reduced in the periwound area at 3 dpi compared to vehicle-treated controls (Fig. 2f–h and Supplementary Fig. 11a). However, at 7 dpi and 14 dpi, when macrophage numbers rebounded (Supplementary Fig. 1m–p), FCER1G+ PDGFRα+ mesenchymal cells reappeared in the wounds (Supplementary Fig. 11b–d).

To determine whether macrophages directly induce the emergence of FCER1G⁺ PDGFRα⁺ mesenchymal cells, we performed an in vitro co-culture assay using NIH/3T3 mouse dermal fibroblasts (MDFs) and bone marrow–derived macrophages isolated from MaFIA mice. MDFs were labeled with CellTrace™ Violet (CTV) and co-cultured with macrophages at a 1:1 ratio for 24 h. Flow cytometric analysis of CTV⁺ MDFs revealed the appearance of a distinct FCER1G⁺ PDGFRα⁺ subpopulation in the co-culture but not in MDFs cultured alone (Fig. 2i–k). Quantification confirmed a significant increase in FCER1G⁺ PDGFRα⁺ mesenchymal cells upon macrophage co-culture, indicating that macrophages directly promote the induction of this mesenchymal subset.

Targeted ablation of Fcer1g-expressing mesenchymal cells leads to less fibrosis
To examine the role of Fcer1g-expressing mesenchymal cells in wound healing, we used STOCK Igs7tm2(tetO-tdTomato,-DTA*G128D)Rdiez/AljJ (Dragon-DTA) mouse. The Dragon-DTA mouse allows doxycycline (Dox)-dependent and recombinase-activated expression of diphtheria toxin A (DTA), leading to cell death37. We crossed Dragon-DTA with C57BL/6-Pdgfraem1(rtTA)Xsun/J (Pdgfra-rtTA) and C57BL/6J-Tg(Fcer1g-cre)NIDA168Htz/Mmucd (Fcer1g-Cre) to generate Pdgfra-rtTA; Fcer1g-Cre; Dragon-DTA mouse (Fig. 3a). In this mouse, only cells co-expressing Pdgfrα and Fcer1g undergo DTA-mediated ablation upon Dox administration (Supplementary Fig. 12a). To achieve targeted ablation, Dox was administered through supplemented food and intraperitoneal injections with optimized dosage and regime (Supplementary Fig. 12b, c) to Pdgfra-rtTA; Fcer1g-Cre; Dragon-DTA (hereafter referred to as “depleted” group) and their control littermates. At 7 dpi, at least 90% of FCER1G+ PDGFRα+ mesenchymal cells were ablated in the wound bed of the depleted group, while FCER1G- PDGFRα+ cells were unaffected (Fig. 3b, c). Immunofluorescence staining for smooth muscle actin (αSMA) at 7 dpi revealed no significant difference between depleted and control wounds (Supplementary Fig. 12d, e), confirming that SMA+ mesenchymal cells were unaffected by the depletion of FCER1G+ PDGFRα+ mesenchymal cells. Macrophage numbers remained unchanged following FCER1G+ PDGFRα+ mesenchymal cell depletion (Supplementary Fig. 12f, g), suggesting a unidirectional regulation in which macrophages regulate this cell population without reciprocal feedback.

We then examined the wound healing process and outcomes in control and depleted groups. At 3 dpi, immunofluorescence analysis demonstrated comparable levels of F4/80⁺ macrophages (Supplementary Fig. 13a, b), CD31⁺ endothelial cells (Supplementary Fig. 13c, d), and K10⁺ keratinocytes between groups (Supplementary Fig. 13g, h), indicating normal inflammatory, angiogenic, and reepithelialization responses. In contrast, PDGFRα staining was reduced in depleted wounds at this early stage (Supplementary Fig. 13e, f), consistent with loss of the Fcer1g-expressing mesenchymal subset. Wound size measurements indicated that wound closure rates were comparable between the control and depletion groups from 0 to 15 dpi (Fig. 3d, e). However, depleted wounds exhibited significantly smaller scar sizes at 21 dpi (Fig. 3d, f), persisting through 8 weeks post-injury (wpi) (Fig. 3d, g and Supplementary Fig. 14a). Hematoxylin and eosin (H&E) staining showed similar healing kinetics between groups, but markedly reduced fibrotic tissue in the depleted groups at 21 dpi (Fig. 3h). Moreover, increased hair follicle formation was observed in the depleted wounds at 8 wpi, suggesting a more regenerative healing outcome. To further quantify the fibrotic outcome, we performed histomorphometric analysis of Masson’s trichrome-stained sections to calculate the scar index (scar area normalized to average dermal thickness) (Supplementary Fig. 14b, c). In 21 dpi wounds, Fcer1g-expressing mesenchymal cell depletion reduced the scar index by over 20% (Fig. 3i), indicating attenuated fibrosis and improved tissue remodeling. Furthermore, total collagen quantification of 21 dpi wounds revealed a 30% reduction in collagen content in the depleted group (Fig. 3j).

To examine the ECM organization, we applied an image-processing algorithm38–40 to profile picrosirius red-stained histological sections from unwounded skin and 8 wpi wounds in both the control and depleted group. Picrosirius red-stained images were deconvoluted into ECM parameters, including color composition (red for mature fibers, green for immature fibers) and fiber properties (length, width, persistence, alignment, etc.). These parameters were quantified independently and analyzed using t-SNE to assess ECM network properties. This analysis revealed that ECM organization in control wounds deviated significantly from that of unwounded skin, with minimal overlap in principal component (PCA) analysis (Fig. 3k). In contrast, ECM organization in the depleted group showed substantial overlap with unwounded skin, indicating reduced fibrosis and a more regenerative matrix structure.

We confirmed that both the control and FCER1G⁺ PDGFRα⁺ mesenchymal cell–depleted groups were in the telogen phase of the hair cycle, ruling out potential confounding effects from differences in hair cycle stage. This was supported by the absence of Ki67 immunostaining in hair follicles (Supplementary Fig. 15a, b) and the lack of nuclear β-catenin staining throughout the experimental timeline (Supplementary Figs. 5a and 16).

Fcer1g-expressing mesenchymal cells are highly proliferative
Previous results revealed a significant increase in Fcer1g-expressing mesenchymal cells following injury (Fig. 2d, e). To determine whether this expansion was driven by proliferation, we administered EdU intraperitoneally to wounded C57BL/6J mice and assessed EdU incorporation in dermal mesenchymal cells 18 hours later. At 3 dpi, Fcer1g-expressing mesenchymal cells exhibited significantly higher EdU incorporation compared to other mesenchymal populations (Fig. 4a–c). These findings indicate that this cell population is highly proliferative during the early phase of wound healing, likely driving its expansion following injury. To examine whether migration also contributed to this expansion, we performed Boyden Chamber assays to evaluate the migratory ability of Fcer1g-expressing mesenchymal cells. The results showed that Fcer1g-expressing mesenchymal cells had significantly lower migratory ability than other mesenchymal cells (Fig. 4d, e). Collectively, these findings indicate that Fcer1g-expressing mesenchymal cells are highly proliferative after injury, and the expansion of Fcer1g-expressing mesenchymal cells is primarily driven by proliferation rather than migration.

EDA fibronectin-deficient wounds have significantly fewer Fcer1g-expressing mesenchymal cells
We next investigated how macrophages induce and regulate Fcer1g-expressing mesenchymal cells. Fcer1g-expressing mesenchymal cells were first mapped onto the comprehensive UMAP, where they clustered as a distinct subpopulation, designated as cluster 28 (Fig. 5f and Supplementary Data file 1). To assess interaction between macrophages (clusters 0, 1, 11, and 20) and Fcer1g-expressing mesenchymal cells (cluster 28), we performed CellChat analysis41. This analysis identified FN1-SDC4 as the top ligand-receptor pair with the highest communication probability between macrophages and Fcer1g-expressing mesenchymal cells at 3 dpi (Supplementary Fig. 17b). Fibronectin (FN1) is known to modulate fibrosis by regulating cell adhesion, migration, differentiation, and intracellular pathways such as TGF-β and β-catenin signaling42–45. In our scRNA-seq dataset, Fn1 was predominantly expressed in macrophages (clusters 0, 1, and 11) and mesenchymal cells (clusters 4 and 9) (Supplementary Fig. 17c). Macrophage depletion significantly reduced Fn1 expression in 3 dpi wounds (Supplementary Fig. 17d), consistent with previous findings that macrophages are a primary source of fibronectin during early wound healing46. These results support a mechanism by which macrophages regulate Fcer1g-expressing mesenchymal cells via fibronectin.

The EDA splice variant of fibronectin is identified as a wound-specific isoform with altered integrin-binding properties in postnatal mice47. Loss of the EDA domain reduces fibrosis, tensile strength, and mesenchymal cell recruitment in wounds45,48. To study whether fibronectin can induce Fcer1g-expressing mesenchymal cells in vivo, we examined 8 dpi wounds in genetically modified B6.129-Fn1tm2Feb mice49, which lack the EDA splice variant of fibronectin (Eda Fn-/-), and their wild-type littermates (Eda Fn+/+). While the number of FCER1G⁻ PDGFRα⁺ cells was comparable between the two groups (Supplementary Fig. 17e), FCER1G⁺ PDGFRα⁺ mesenchymal cells were significantly reduced in Eda Fn-/- wounds (Supplementary Fig. 17f-h).

Fcer1g-expressing mesenchymal cells are conserved in human wound healing
To test whether FCER1G⁺ PDGFRα⁺ mesenchymal cells are conserved in human wound healing, we analyzed a previously published human skin wound healing scRNA-seq dataset (GSE241132) collected at days 0, 1, 7, and 30 post-injury50. Within this dataset, PDGFRA and PDGFRB marked mesenchymal populations (Supplementary Fig. 18a, b), which were further subclustered into 13 distinct subclusters (Fig. 5a). Using gene signatures derived from mouse Fcer1g-expressing mesenchymal cells, as well as FCER1G expression alone, we identified a human mesenchymal subcluster (cluster 11) that uniquely expressed both the mouse Fcer1g signature (Fig. 5b, c) and FCER1G itself (Supplementary Fig. 18c, d). The abundance of human FCER1G-expressing mesenchymal cells (cluster 11) peaked at day 7 post-injury (Fig. 5d, e), consistent with the kinetics observed in the mouse model (Supplementary Fig. 10b). FCER1G also exhibited one of the highest U-scores in human mesenchymal subcluster 11 (Supplementary Fig. 18e and Supplementary Data file 2). Moreover, the top differentially expressed genes (DEGs) of FCER1G⁺ PDGFRα⁺ mesenchymal cells were highly conserved between human and mouse, suggesting this represents an evolutionarily conserved mesenchymal population that responds to skin injury (Fig. 5f and Supplementary Data file 3).

To evaluate whether FCER1G⁺ PDGFRα⁺ mesenchymal cells are associated with human scar formation, we further reanalyzed another publicly available bulk RNA-seq dataset (GSE178411), which includes 108 human samples across uninjured skin (n=26), early wounds (n=22), late wounds (n=29), chronic wounds (n=3), and hypertrophic scars (n=28). The top 100 DEGs derived from both mouse and human FCER1G⁺ PDGFRα⁺ mesenchymal cells effectively distinguished wound and scar samples from uninjured skin (Fig. 5g and Supplementary Fig. 18f). ssGSEA analysis revealed that FCER1G-associated signatures peaked during the early (acute) wound phase and gradually declined as wounds progressed toward hypertrophic scarring (Supplementary Fig. 19a), consistent with the temporal dynamics observed in mice. Furthermore, FCER1G expression alone recapitulated this pattern (Supplementary Fig. 19b), supporting its role as a key marker of a pro-fibrotic mesenchymal subset involved in hypertrophic scar formation during wound healing.

Methods
Mice
We obtained institutional ethical approval through Duke University Institutional Animal Care and Use Committee (IACUC) of Duke University for all animal experiments under the protocol number A1512307. Young adult mice (8 weeks of age) in both sexes were used in this study. C57BL/6J, MaFIA (C57BL/6-Tg(Csf1r-EGFP-NGFR/FKBP1A/TNFRSF6)2Bck/J, Strain# 005070)18, Pdgfra-rtTA (C57BL/6-Pdgfraem1(rtTA)Xsun/J, Strain# 034459)66 and Dragon-DTA (STOCK Igs7tm2(tetO-tdTomato,-DTA*G128D)Rdiez/AljJ, Strain# 034778)37 were purchased from The Jackson Laboratory. Fcer1g-Cre (C57BL/6J-Tg(Fcer1g-cre)NIDA168Htz/Mmucd, RRID: MMRRC_043798-UCD)67 was obtained from the Mutant Mouse Resource and Research Center at the University of California at Davis. Both male and female mice were included in all experiments; however, the study was not powered for sex-specific analysis, and sex differences were not a predefined endpoint. Mice were housed under conditions at 22°C, with a humidity between 30% to 70%, and a light cycle of 12 h-12 h on-offset. Primers used for genotyping of genetically engineered mouse models were included in Supplementary Table 2.

Macrophage depletion
To conditionally deplete macrophages, we used the MaFIA mouse. AP20187 (Takara, Cat#532522) was dissolved in 100% ethanol to a concentration of 62.5 mg/mL and stored at −20°C. This stock solution was further diluted to a 2.5 mg/mL in an injection solution of 10% PEG-400, and 2% Tween-20 in DI water. Retro-orbital injection of AP20187 at a dose of 10 mg per kg body weight or vehicle were used to ablate macrophages. Five days of treatment effectively depletes 90% of macrophage cells in the bone marrow68.

Depletion of Fcer1g-expressing mesenchymal cells
To deplete Fcer1g-expressing mesenchymal cells in the skin, we generated Pdgfra-rtTA; Fcer1g-Cre; Dragon-DTA mouse. As Pdgfra-rtTA mice were firstly reported to target lung mesenchymal cells66, we evaluated their efficiency in targeting dermal mesenchymal cells. Dragon-DTA includes a tdTomato (tdTM) reporter, allowing visualization of cells targeted by the rtTA system. In Pdgfra-rtTA; Dragon-DTA mice, Dox administration induced tdTM+ cells in the wounded skin (Supplementary Fig. 12c). The highest targeting efficiency was achieved by combining Dox-supplemented food (625 mg/kg diet) and intraperitoneal injections (100 mg/kg body weight). At this dosage, over 80% of tdTM+ cells exhibited PDGFRα immunoreactivity, confirming efficient targeting of dermal PDGFRα+ mesenchymal cells in Pdgfra-rtTA mice. We then followed the optimized dosage and regimen (Supplementary Fig. 12b) to administrate Dox to Pdgfra-rtTA; Fcer1g-Cre; Dragon-DTA and their control littermates.

Excisional wound splinting model
The excisional wound splinting model was conducted according to an established protocol69. Mice were anesthetized with isoflurane, hairs were clipped, the skin site was disinfected and two full-thickness excisional wounds on the dorsal skin was created with a sterile 5 mm biopsy punch. Sterilized silicone splints (14 mm/7 mm) were centered over the wounds and anchored with nylon sutures. The wounds were then covered with a transparent occlusive dressing (Tegaderm) to maintain moisture, and the mouse was wrapped in Vetwrap before being placed in a single cage.

Cell dissociation
Mice were euthanized by CO2 exposure followed by cervical dislocation. The dorsal skin was disinfected, shaved, depilated, and excised. The excised skin was cut into small pieces (approximately 0.2 cm × 0.2 cm) and transferred into a 15 mL conical tube. Each skin piece was incubated in 5 mL of dissociation buffer at 37°C for 60 min. The dissociation buffer contained 5 mL RPMI 1640 (no calcium or magnesium) with 232 U DNAse I, 0.25 mg/mL Libera stock (2.5 mg/mL), 23.2 mM HEPES, and 2.32 mM sodium pyruvate. The digestion was stopped by adding 10 µL of 0.5 M EDTA and 400 µL FBS, and the samples were transferred to GentleMacs tubes (Miltenyi Biotec, Cat#130-093-237) for dissociation (program: m_impTumor_01.01).

The digested samples were filtered through a 70 µm filter into a 50 mL conical tube, rinsed with 10 mL of RPMI media containing FBS, and filtered again through a fresh 70 µm filter. The samples were centrifuged at 500 × g for 10 min at 4°C, and the supernatant was carefully aspirated. The cell pellets were resuspended in flow cytometry buffer, which was prepared using 500 mL PBS, 2 mL of 0.5 M EDTA, 25 mL FBS, and 5 mL non-essential amino acids (100x stock).

scRNA sequencing of mouse model
Three MaFIA mice per condition were studied. Cell dissociation was prepared as above, and then filtered through 30 μm cell strainer. Propidium iodide (PI) was added to each sample before loading to a DiVa cell sorter (BD Biosciences) for flow cytometry analysis or cell sorting. 18 K live cells were sorted and loaded on the 10x Genomics Chromium using the Chromium Single Cell 3’ Reagent V3 Kit, and the sequencing libraries were constructed following the user guide. Cell Ranger version V3.0.2 (10x Genomics) was used to process raw sequencing data before subsequent analyses. These RNA sequencing reads were then aligned against the refdata-cellranger-mm10-3.0.0 transcriptome to quantify the expression of transcripts in each cell to create feature-barcode matrices. There were around 12 K cells recovered per sample. The median genes per cell were around 2200. The analyses of processed scRNA-seq data were carried out in R version 4.4.0 using the Seurat v4 for downstream analysis31,32.

To eliminate empty droplets, doublets, and dead cells, as well as cells with high mitochondrial DNA expression, we applied a filtering criterion of 200 < nFeature < 2500, percent.mt < 5 and doublets were removed by DoubletFinder70. During preprocessing, we also tested different thresholds for the number of features, setting them at 1800 and 7000 based on density plots (Supplementary Fig. 6a), which corresponded to the first enriched peak and the entire cell population, respectively. However, these thresholds did not provide a clear and distinct separation between clusters in the downstream analysis when compared to the 2500 threshold.

We then normalized, anchored and integrated the filtered cells and performed the non-linear dimensional reduction by UMAP. We selected mesenchymal cell clusters by canonical mesenchymal markers, including Col1a1, Col1a2, Pdgfrα, Col5a1, Loxl1, Lum, Fbln1 and Cd34. We then re-anchored and re-integrated to define the mesenchymal subpopulations. Differentially expressed genes with top 100 log2FC from the FindMarkers function were used as the signature genes in ssGSEA.

CellChat was used to analyze and visualize the cell-cell communication network41. In general, the unique Fcer1g-expressing mesenchymal cell population was first mapped to the comprehensive UMAP as cluster 28 (Supplementary Fig. 17a), which was saved and loaded as a new scRNA-seq matrix for further analysis. Clusters 0, 1, 11, 20 and 28 from the vehicle group were extracted to analyze the interaction between macrophages and Fcer1g-expressing mesenchymal cells.

scRNA sequencing re-analysis of publicly available cohort
Data were obtained from GSE241132. Analyses were performed in Seurat (v5.3.0) following the user guide. Briefly, low-quality cells were filtered out, and expression values were normalized. Datasets were then integrated using an anchor-based approach. Dimensionality reduction was performed with Uniform Manifold Approximation and Projection (UMAP), and cells were clustered using a shared-nearest-neighbor (SNN) modularity-optimization algorithm. Clusters were annotated based on marker-gene expression. After extracting the fibroblast cluster from the full dataset, dimensionality reduction and clustering were repeated as above. Cell-cycle scores (S and G2M) were computed from cell-cycle–related gene expression. Differential expression analysis was performed with the FindMarkers function, comparing the FCER1G-high cluster with resting clusters. Differentially expressed genes with top 100 log2FC from the FindMarkers function were used as the signature genes in ssGSEA.

Bulk RNA sequencing re-analysis of publicly available cohort
A bulk RNA-seq transcriptomic dataset for human acute skin wound were downloaded from GSE178411. The downloaded FASTQ files were aligned to GRCh38 reference genome using STAR71, and the raw gene counts were computed as default –quantMode. Following normalization and differentially expressed gene calling was performed using the DESeq2 R package72 with default parameters. VST normalization of DESeq2 counts were performed prior to assessing single gene expression levels. ssGSEA with the human or mouse FCER1G+ signature was performed using the default parameters, with an alpha value equal to 0.25. NES enrichment score was calculated based on GSEA with 10,000 permutations, which is shown by the running sum statistics plot.

Histological analysis
Wound tissues were harvested using a 7 mm biopsy punch at designated time points after complete wound closure, ensuring capture of the entire wound bed. Each wound sample was bisected through the center, fixed immediately in ice-cold 4% paraformaldehyde (PFA) for 36 h under gentle agitation at 4°C, processed and embedded such that cross-sections were oriented with the wound center facing downward. Serial sections were then collected in 5 μm and deparaffined before subject to staining. Tissues from B6.129-Fn1tm2Feb mice (Eda Fn-/-) and their wild-type littermates (Eda Fn+/+) were collected and prepared following an established procedure45. Serial sections through the circular wound were examined to identify the center, and the staining was performed on the wound center to ensure fair comparison.

For Masson’s trichrome staining, we followed the EMS technical Data sheet (Cat#26367-series). In brief, slides were mordanted in Bouin’s Fixative for 1 h at 56°C. After washing, they were stained with Weigert’s Iron Hematoxylin for 5 min and washed in distilled water for 10 min. Sections were then stained in Biebrich Scarlet-Acid Fuchsin for 15 minutes, followed by treatment with Phosphomolybdic Acid-Phosphotungstic Acid for 15 min. Aniline Blue was applied for 10 to 20 min, and differentiation was performed using 1% Acetic Acid for 3 to 5 min. For picrosirius red staining, we followed the Abcam protocol (Cat#ab150681). In brief, slides were mordanted in picrosirius red solution for 1 h and rinsed quickly with 2 changes of 0.5% acetic acid. Finally, tissues were dehydrated in alcohol, cleared in xylene, and mounted.

For immunofluorescence staining, slides were steamed with 1% citrate-based antigen retrieval (Vector Laboratories, H-3300) for 10 mins and blocked with 5% donkey serum and 0.3% Triton-X-100 in 1x PBS overnight at 4°C prior to addition of the following primary antibodies: Rat-F4/80 (Biorad, Cat#MCA497GA, clone A3-1, 1:500), Rabbit-Fcer1g (Invitrogen, Cat#PA5-115222, 1:200), Goat-Pdgfra (R&D, Cat#AF1062; 1:100), Rabbit-Keratin 10 (BioLegend, Cat# 905403, 1:100), Rat-CD31 (BDBiosciences, Cat#557355, clone MEC13.3, 1:100), Rabbit-Ki67 (Novus, Cat#NB500, 1:1000) and Rabbit-alpha smooth muscle actin (Proteintech Cat#80008-1-RR, Clone No. 5H7 1:500). Slides were steamed with 1% Citrate-based antigen retrieval (Vector Laboratories, H-3300) for 10 mins, incubated with 4% PFA for 8 mins to increase permeabilization, and blocked with 3% bovine serum albumin, 0.5% Donkey serum in 1x TBST for 1 h at room temperature prior to Chicken-GFP (Abcam, cat# ab13970, 1:100) primary antibody. Rat-IgG (Invitrogen, Cat#02-9602), Rabbit-IgG (R&D, Cat#AB105-C), Goat-IgG (R&D, Cat#AB108-C) and Chicken-IgY (R&D, Cat#AB101-C) were used at corresponding concentrations as negative controls. Slides were then incubated for 1 h with Goat anti-rat Alexa Fluor 647 (Invitrogen, Cat#A-21247, 1:500), Goat anti-rat Alexa Fluor 594 (BioLegend, Cat#A-405422, 1:500), Donkey anti-goat Alexa Fluor 647 (Abcam, Cat#ab150131, 1:500), Goat anti-rabbit Alexa Fluor 594 (Invitrogen, Cat#A-11037, 1:500), Goat anti-chicken (Invitrogen, Cat#A-11039, 1:500), Goat anti-rabbit Alexa Fluor 647 (Invitrogen, Cat#A-32733, 1:500). Finally, slides were mounted in ProLong™ Glass Antifade Mountant with NucBlue™ Stain (Invitrogen, Cat#P36981).

For immunohistochemistry, slides were steamed with 1% citrate-based antigen retrieval (Vector Laboratories, H-3300) for 15 mins, blocked with 3% H₂O₂ in methanol for 10 mins, and subsequently blocked in 1× PBST containing 1.5% normal goat serum for 60 mins prior to following primary antibodies: Rabbit-Fcer1g (Invitrogen, Cat#PA5-115222, 1:200) and Rabbit-beta Catenin (Invitrogen, Cat#71-2700, 1:400). Rabbit-IgG (R&D, Cat#AB105-C) were used at corresponding concentrations as negative controls. Slides were then incubated overnight at 4°C in a humidified chamber. The next day, slides were equilibrated to room temperature, washed with PBST, and incubated with biotinylated secondary antibody for 30 min, followed by Vectastain ABC reagent for an additional 30 min. After PBST and water washes, signals were developed using Vector ImmPACT DAB, with color development monitored microscopically. Reactions were stopped in water, and sections were counterstained with hematoxylin, rinsed, and briefly incubated in PBS. Slides were then dehydrated through graded ethanol, cleared, air-dried, and mounted with Cytoseal.

Brightfield images were acquired with a Leica DM2000 LED microscope, while fluorescent images were acquired with a Leica DM6000 SP5 upright confocal microscope and a Leica Stellaries 8 confocal microscope. Images were processed in Fiji (ImageJ) using identical settings across experimental groups; no nonlinear adjustments were applied.

Measuring cell proliferation in vivo
To assess cell proliferation, EdU (Invitrogen: A10044) was injected intraperitoneally at 25 mg/kg body weight into mice 18 hours before euthanasia. Cells were dissociated following the procedures as described earlier in the Cell Dissociation section. We followed the protocol from the Click-iT™ EdU Flow Cytometry Assay Kit (Invitrogen, Cat# MP 35002) for cell permeabilization and EdU detection. Finally, cells were analyzed by flow cytometry.

Cell culture
For the Boyden chamber assay to assess cell migration, mesenchymal cells were sorted from 3 dpi wound and were seeded at 4000 cells per 8 µm 24-well insert (Falcon, Cat#353097) with serum-free medium, while the bottom wells were filled with medium with FBS. After 2 days, the inserts were collected, washed with PBS, and fixed with 4% PFA for 2 min. Cells were permeabilized with 100% methanol for 20 minutes and stained with 0.1% crystal violet (20% methanol in PBS). Non-migrated cells were scraped off, and the membrane was imaged.

To differentiate macrophages from bone marrow stromal cells, mice were euthanized by CO2 exposure followed by cervical dislocation. The femur and tibia were dissected, and the ends of the bones were removed. The bones were placed cut-side down in a 0.6 mL tube with a hole at the bottom, inside a 1.5 mL Eppendorf tube. Bone marrow was extracted by centrifugation at 10,000 x g, resuspended in 5 mL of plating medium (10% FBS, 1% Pen/Strep in alpha-MEM), and centrifuged at 500 x g for 5 minutes. After discarding the medium, 1 mL of red blood cell lysis buffer was added for 1 min at room temperature. The cells were washed with 5 mL of plating medium, centrifuged again, and resuspended in 10 mL of fresh medium. The cell suspension was filtered through a 70 µm sterile cell strainer, plated in a 10 cm dish with 40 ng/mL M-CSF (R&D, Cat#416-ML), and cultured for 5 days. Macrophage identity was confirmed by F4/80 and CD11b staining.

NIH/3T3 murine dermal fibroblasts (MDFs) were maintained in DMEM supplemented with 10% FBS and 1% penicillin–streptomycin at 37°C in a humidified incubator with 5% CO₂. Cells were passaged every 2–3 days at 70–80% confluence. For passaging, cultures were washed with PBS and incubated with 0.05% trypsin–EDTA for 5 min at 37°C. Trypsinization was quenched by adding 5 mL complete medium, and cells were collected by centrifugation at 500 × g for 5 min before resuspension and replating.

For co-culture experiments, macrophages were dissociated with 0.25% trypsin-EDTA on day 5, counted, and replated with mesenchymal cells. NIH/3t3 MDF were counted and stained with CTV dye (1 µL of 5 mM CTV for 1 million cells in 1 mL) from the CellTrace™ Violet Cell Proliferation Kit (Invitrogen, Cat#C34557) for 20 minutes at 37°C in the dark. After washing, MDF were co-cultured with macrophages at a 1:1 ratio (150 K cells each) or plated alone (300 K cells) in a 6-well plate for 24 h. Afterward, cells were dissociated and analyzed by flow cytometry.

Flow cytometry
Skin cells were dissociated as described earlier in the Cell Dissociation section and incubated with the antibodies for 30 min at room temperature, protected from light. Antibodies used in this study include PDGFRA-APC (Invitrogen, Cat#17-1401-81, 1:50), FCER1G-PE (Miltenyi Biotec, Cat#130-118-760, 1:200), F4/80-AF647 (BDBiosciences, Cat#565853, 1:100) and CD11b-PE/Cy7 (BioLegend, Cat#101216, Clone# M1/70, 1:100). After incubation, cells were washed twice with 1x PBS and filtered through a 30 µm filter. Flow cytometry was then performed using a FACSCanto II flow cytometer (BDBiosciences), and data were analyzed with FlowJo v.10 (TreeStar). For cell sorting, a DiVa cell sorter (BD Biosciences) was used. A representative gating strategy is provided in Supplementary Fig. 20.

Total collagen quantification
We followed the protocol from the Total collagen assay kit (Abcam, Cat#ab222942) to quantify the total collagen in the wounded skins. In brief, minced tissues were hydrolyzed with 10 N NaOH at 120°C for 3 h to yield hydroxyproline, which was further oxidized and developed. The absorbance was detected by SpectraMax i3x plate reader (Molecular Devices) at OD 560 nm and compared to standards to calculate the final concentration.

Morphometric analysis
The macroscopic wound area was quantified by processing of photographs taken at various time points and was calculated as the percentage of the wound area immediately post-surgery. The scar index was used to assess the extent of scar formation by calculating the ratio of the contoured area of the scar (from a sagittal view) to the average thickness of normal skin.

Quantitative analysis of ECM organization
Skin sections that were 60 µm from the center of the wound were stained with picrosirius red as described above. Wounded skin and surrounding unwounded skin area were imaged for a minimum of 18 images per experiment condition and subject for quantitative analysis of ECM network properties38,39. In general, picrosirius Red-stained images was first deconvoluted to isolate different color contributions corresponding to mature and immature connective tissue fibers, excluding cellular elements. The deconvoluted images are then processed with an adaptive Wiener filter to reduce noise, followed by binarization and morphological filtering to extract fiber-shaped objects. The fiber network is skeletonized, and various properties like fiber length, width, and alignment are measured using the regionprops function. Finally, dimensionality reduction of the quantified data is performed using t-SNE to visualize group differences.

Statistical analysis
Data are presented as mean ± standard deviation (s.d.), with individual values shown for each sample. Statistical analysis of scRNA-seq cell distribution percentages was conducted using a two-way repeated measures ANOVA in JMP Pro 16 with the Full Factorial Repeated Measures ANOVA Add-In. Other statistical analyses were performed using GraphPad Prism 10, applying unpaired or paired two-tails t tests as appropriate, following confirmation of normal distribution.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript addresses the role of macrophage-fibroblast interactions during skin repair after injury. The authors utilize MAFIA mice to specifically deplete Csfr1a+ macrophages after injury and identify a population of mesenchymal cells that is reduced in MAFIA mice 3 days after injury. The authors identify Fcer1g as differentially expressed in this mesenchymal subpopulation. Using mouse genetics, the authors deplete Pdgfra+Fcer1g+ cells and find that wounds do not heal as in control. The authors then analyze FN1 expression by macrophages as a regulator of Fcer1g expressing fibroblasts using B6.129-Fn1tm2Feb mice and find a reduction in these cells in wounds of these mice. Overall, this paper has some interesting findings but lacks a robust characterization of the mouse models to fully support the conclusions drawn.\nSpecific concerns:\n1. Wounding protocol/analysis:\nA. The images shown indicate that the skin of the mice may not be in the same stage of the hair cycle, which can influence wound healing and cell heterogeneity (see D. M. Ansell et al. Journal of Investigative Dermatology (2011) and references therein). The authors should ensure that the wounds and analyses were performed in the same hair cycle stage. Specifically Figs. 1d and S3f look to be in different stages.\nB. The analysis of the wounds of the mice do not appear to be not consistently in the center of the wounds. Why would the wounds without macrophages have smaller wound areas if they are not healing as effectively? What is the reepithelialization and angiogenesis process in the authors' wound paradigm in control and MAFIA mice? I suspect that the smaller wound size is due to analysis of wounds at the edge rather than the center of the wound.\n2. Macrophage depletion and analysis: The description of the experimental paradigm indicates that the authors induced depletion of Csfr1a+ cells for 5 days PRIOR to wounding, which would likely deplete tissue resident macrophages rather than myeloid-derived macrophages, which are the prevalent macrophages recruited to skin wounds after injury. The authors should demonstrate with an unbiased approach -- like FACS analysis of macrophages how the system impacts macrophages especially 3 days post injury. Ideally, the authors should analyze which populations of macrophages (not just GFP+ cells) are impacted in their experimental system. The scRNA sequencing results in S1f and S1e suggest a reduction but this technique is not a good system for determining cell loss alone without other experimental systems.\n3. Validation of Fcer1g in wound of MAFIA mice 3 days after injury.\nA. While the authors show that Fcer1g+ cells are reduced adjacent form the wound, the authors only validate the reduction of Fcer1g+ cells after injury by FACS rather than immunostaining. Is there a reduction in the wound bed?\nB. Fig. 2c lacks green or red cells as indicated in the figure label.\nC. The Fcer1g+ cells appear to be around larger organized structures like blood vessels -- did the authors analyze whether they colocalize with CD31?\n4. Depletion of Fcer1g+ cells.\nA. Since Fcer1g seems to be expressed in a subset of mesenchymal cells, why are all of the Pdgfra+ cells depleted in Fig.3b. The authors should analyze the other types of mesenchymal cells that might remain in the skin after depletion--are there any SMA+ cells?\nB. Is Fcer1g expressed in other cell types in the skin?\nC. The authors should characterize wounds of the Fcer1g depletion mouse model prior to D7-- is inflammation normal? How does epithelialization, revascularization, fibroblast repopulation occur after injury?\n5. I am not sure the deletion of FN1 adds much to the paper given the lack of specific depletion in macrophages and the poor characterization of the model in the paper. The immunostaining in Fig. 5 is not very convincing and the quantification is not consistent with the prior analysis.\nMinor comments:\n1. It would helpful to include the annotation of the cell types from Fig. 1F in Fig. S1.", "reviewer_2": "The paper explores a novel mesenchymal cell subpopulation activated by macrophages that induces fibrosis. It would be beneficial to mention whether this subpopulation exists in human skin, as the potential for this to translate into a therapy remains unclear.\nComments\n1. Validating FCER1G expression in human wound edge data (e.g., GSE178411) would be helpful.\n2. The Macrophage depletion method section (Line 319-325) requires clarification. It mentions retro-orbital injection of AP20187; however, the original reference describes intraperitoneal injections. Please confirm and correct the method used for macrophage depletion.\n3. The study's major limitation is its lack of validation using human data or in vitro experiments.\n4. The Cell Culture Methods section (Lines 438–450) requires clarification. It states that mouse tails were used to isolate primary dermal mesenchymal cells. However, given the focus of the study, it would be more appropriate to isolate these cells from the wound edge tissue and perform the cell proliferation assay accordingly. If tail tissue was intentionally used, please justify this choice.\n5. In general, few connections appear to be made regarding the potential therapeutic effects of diminishing the subpopulation. It would also be interesting to see if increasing the subpopulation has any impact. Additionally, there is no in vitro or human data to support the in vivo work, which could better demonstrate the link with fibronectin.", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "We thank the reviewers for their thoughtful evaluation and recognition of the novelty of our work. We have revised the manuscript to include additional analyses characterizing the mouse model in both the Result and Discussion sections, as detailed in our responses to the reviewers’ comments.\nSpecific concerns:\n1. Wounding protocol/analysis:\nA. The images shown indicate that the skin of the mice may not be in the same stage of the hair cycle, which can influence wound healing and cell heterogeneity (see D. M. Ansell et al. Journal of Investigative Dermatology (2011) and references therein). The authors should ensure that the wounds and analyses were performed in the same hair cycle stage. Specifically Figs. 1d and S3f look to be in different stages.\nWe agree with the reviewer about the importance of hair cycle synchronization, which is indeed a critical factor influencing wound healing. We took several steps to ensure consistency across all experiments.\nAll animals used in this study were on a C57BL/6J background and were strictly age- and sex-matched. According to established literature (Müller-Röver et al., J. Invest. Dermatol., 2001), the dorsal skin of C57BL/6J mice remains in the telogen phase from approximately 7 to 12 weeks of age (Supplementary Figure 3a). Accordingly, all wounding procedures were performed at 8 weeks of age, ensuring that skin injury occurred during the telogen phase. Since all experimental groups were wounded at the same age, tissue collection and analysis time points were also synchronized between groups. This design ensured that both wound induction and subsequent analyses were conducted within the same phase of the hair cycle across all animals.\nTo further validate this, we performed immunohistochemistry for Ki67 and nuclear β-catenin—established markers of the anagen phase (Magerl et al., J. Invest. Dermatol., 2001; DasGupta and Fuchs, Development, 1999). Both markers were absent from hair follicles in MaFIA mice treated with either vehicle or AP20187 (Supplementary Figure 3), as well as in Pdgfra-rtTA; Fcer1g-Cre; Dragon-DTA mice and their littermate controls (Supplementary Figure 7). These results confirm that all mice remained in the telogen phase throughout the duration of the study.\nWe have added the following text to the revised manuscript to clarify these points:\nResults, page 3, lines 107–121:\n“Previous studies have shown that hair follicles in the anagen phase can actively contribute to mouse wound re-epithelialization [21], making it critical to ensure consistent hair cycle phases across experimental groups. The anagen phase is characterized by active cell proliferation [22], as indicated by robust Ki67 expression (Supplementary Figure 3b), and by activated Wnt/β-catenin signaling [23], evidenced by nuclear β-catenin localization (Supplementary Figure 3d). Around 7 weeks of age, follicles transition into the telogen phase, marked by the absence of both Ki67 and nuclear β-catenin, and remain in this phase until at least 12 weeks postnatal (Supplementary Figure 3a–b). In our study, mice were wounded at approximately 8 weeks of age and analyzed around 11 weeks, during which time hair follicles are expected to be in the telogen phase. Consistent with this, both vehicle- and AP20187-treated groups showed no follicular Ki67 expression throughout the experimental timeline (Supplementary Figure 3c), indicating limited proliferative activity. Moreover, the absence of nuclear β-catenin in both groups (Supplementary Figure 3e) further confirmed that all animals remained in the telogen phase during the study period.”\nResults, page 7, lines 271–275:\n“We confirmed that both the control and FCER1G⁺ PDGFRα⁺ mesenchymal cell–depleted groups were in the telogen phase of the hair cycle, ruling out potential confounding effects from differences in hair cycle stage. This was supported by the absence of Ki67 immunostaining in hair follicles (Supplementary Figure 7a–b) and the lack of nuclear β-catenin staining throughout the experimental timeline (Supplementary Figures 3d and 7c).”\nTo avoid confusion, we have also replaced the images in Figure 1e with more representative ones that better display hair follicle morphology.\nB. The analysis of the wounds of the mice do not appear to be not consistently in the center of the wounds. Why would the wounds without macrophages have smaller wound areas if they are not healing as effectively? What is the reepithelialization and angiogenesis process in the authors' wound paradigm in control and MAFIA mice? I suspect that the smaller wound size is due to analysis of wounds at the edge rather than the center of the wound.\nWe address each point in detail below:\n1. Regarding the wound sample collection and preparation:\nTo clarify the methodology used for wound tissue collection and histological analysis, we added additional text to the Method section (page 13, lines 541-548):\n“Wound tissues were harvested using a 7-mm biopsy punch at designated time points after complete wound closure, ensuring capture of the entire wound bed. Each wound sample was bisected through the center, fixed immediately in ice-cold 4% paraformaldehyde (PFA) for 36h under gentle agitation at 4 °C, processed and embedded such that cross-sections were oriented with the wound center facing downward. Serial sections were then collected in 5 μm and deparaffined before subject to staining.”\nTo further illustrate this visually, we included serial H&E sections collected at 50 μm intervals from the wound center in Supplementary Figure 2c as supportive data for Figure 1e and added the corresponding explanation in the Results section (page 3, lines 99–102):\n“Serial sections demonstrated reduced fibrosis in AP20187-treated wounds spanning from the wound center to the edge, as fibrosis was nearly absent at 150 μm from the center in the depleted group, while still clearly visible at 250 μm in controls. (Supplementary Figure 2c).”\nThe representative images in Figure 1e were acquired at a consistent depth (50 μm) from the wound center to allow for standardized comparisons. Thus, all histological data presented in the main figures reflect central wound sections rather than peripheral areas. This standardized and centrally focused sectioning approach was applied across all groups to ensure consistency and minimize spatial bias in histological interpretation.\n2. Regarding wound size, healing effectiveness, re-epithelialization, and angiogenesis:\nTo clarify the effects of macrophage depletion on wound healing dynamics, we evaluated both epithelial closure and vascularization in vehicle- and AP20187-treated groups. In the revised manuscript, we quantified the distance between epithelial tips at 7 and 14 days post-injury (dpi) (Supplementary Figure 2d–g) and found no statistically significant difference in wound size at 7 dpi. By 14 dpi, all wounds were fully re-epithelialized in both groups, indicating that macrophage depletion did not impair wound healing effectiveness or re-epithelialization.\nWe also assessed angiogenesis by performing immunofluorescence staining for CD31 to quantify vascular density in the same wounds. At both 7 and 14 dpi, the number of CD31⁺ endothelial cells per high-power field (HPF) was comparable between vehicle- and AP20187-treated wounds (Supplementary Figure 2h–k), suggesting that macrophage depletion did not significantly alter angiogenesis.\nWe added these findings in the Result section as follows (page 3, lines 102–106):\n“Moreover, immunofluorescence staining for keratin 10 (K10) and CD31 revealed no significant differences in reepithelialization or angiogenesis between vehicle- and AP20187-treated wounds at 7 dpi and 14 dpi (Supplementary Figure 2d–k), indicating that the reduction in fibrotic area specifically reflects macrophage-dependent regulation of wound fibrosis.”\n3. Regarding the scar size:\nTo provide a more complete representation of scar morphology, we included macroscopic scar images (Supplementary Figure 2a) and Masson’s trichrome–stained sections (Supplementary Figure 2b) from all vehicle- and AP20187-treated (macrophage-depleted) wounds as supportive data for Figures 1c and 1e, respectively. The scar area quantification shown in Figure 1d is derived from the macroscopic wound areas presented in Figures 1c and Supplementary Figure 2a.\nWe clarified this in the Result section as follows (page 3, lines 95–99):\n“…early-phase macrophage depletion led to significantly reduced fibrosis at 21 dpi compared to vehicle-treated controls (Figure 1c–d, Supplementary Figure 2a). Masson’s trichrome staining revealed that AP20187-treated wounds exhibited smaller areas of thick, densely packed collagen fibers, indicating attenuated fibrosis (Figure 1e, Supplementary Figure 2b).”\nThese additional images and quantifications collectively confirm that the reduction in scar size observed in macrophage-depleted wounds is consistent across both macroscopic and histological analyses.\n2. Macrophage depletion and analysis: The description of the experimental paradigm indicates that the authors induced depletion of Csfr1a+ cells for 5 days PRIOR to wounding, which would likely deplete tissue resident macrophages rather than myeloid-derived macrophages, which are the prevalent macrophages recruited to skin wounds after injury. The authors should demonstrate with an unbiased approach -- like FACS analysis of macrophages how the system impacts macrophages especially 3 days post injury. Ideally, the authors should analyze which populations of macrophages (not just GFP+ cells) are impacted in their experimental system. The scRNA sequencing results in S1f and S1e suggest a reduction but this technique is not a good system for determining cell loss alone without other experimental systems.\nTo address this, we evaluated the CD11b⁺ and F4/80⁺ macrophage populations at 3 dpi using both flow cytometry and immunofluorescence (Supplementary Figure 1a–l) to assess how AP20187 impacts distinct macrophage subsets in MaFIA mice, including dermal resident macrophages and wound-infiltrating myeloid-derived macrophages.\nTo assess the impact on dermal resident macrophages, we analyzed skin macrophages at 3 dpi and observed robust depletion following AP20187 treatment (Supplementary Figure 1i–l). To evaluate the effects on myeloid-derived macrophages, we examined the bone marrow (Supplementary Figure 1a–d) and spleen (Supplementary Figure 1e–h) at the same time point and similarly observed significant depletion. Together, these data demonstrate that the MaFIA system effectively targets both tissue-resident and bone marrow–derived macrophage populations.\nWe added this to the Result section (page 2-3, lines 85–92) as:\n“To further confirm that GFP⁺ depletion reflects loss of macrophages, we performed flow cytometry for F4/80 and CD11b double-positive cells in the bone marrow, spleen, and skin from vehicle- and AP20187-treated MaFIA mice (Supplementary Figure 1). These analyses confirmed robust depletion of F4/80⁺CD11b⁺ macrophages in the skin (>75%) (Supplementary Figure 1i–j) and moderate depletion in the bone marrow and spleen (~50%) (Supplementary Figure 1a–b and 1e–f). Immunofluorescence staining further validated the loss of F4/80⁺ macrophages in these tissues (Supplementary Figure 1c–d, 1g–h, and 1k–l).”\nThese findings are consistent with our scRNA-seq analysis (Supplementary Figure 4d–e) and collectively support the conclusion that AP20187 treatment induces broad and systemic macrophage depletion in this system.\n3. Validation of Fcer1g in wound of MAFIA mice 3 days after injury.\nA. While the authors show that Fcer1g+ cells are reduced adjacent form the wound, the authors only validate the reduction of Fcer1g+ cells after injury by FACS rather than immunostaining. Is there a reduction in the wound bed?\nIn the revised manuscript, Figure 2f–g includes representative immunostaining and quantification demonstrating a reduction of FCER1G⁺ PDGFRα⁺ mesenchymal cells in 3 dpi MaFIA wounds following macrophage depletion. To improve clarity, we have separated the immunofluorescence channels for FCER1G and PDGFRα in Figure 2f, which clearly shows the decrease in FCER1G⁺ PDGFRα⁺ cells upon AP20187 treatment.\nAt 3 dpi, the wound bed contains minimal granulation tissue and sparse PDGFRα⁺ mesenchymal cellularity, making reliable quantification within this area difficult. Therefore, quantification was performed in the periwound region, where mesenchymal cells are consistently present and structurally defined. To provide a broader spatial context, Supplementary Figure 5i shows wide-field immunofluorescence of FCER1G and PDGFRα staining across distances ranging from 10 µm to 2500 µm from the wound edge.\nIn addition, Supplementary Figures 5j–l demonstrate that FCER1G⁺ PDGFRα⁺ mesenchymal cells begin to repopulate the wound area by 7 and 14 dpi as macrophages return, further supporting macrophage-dependent regulation of this mesenchymal population. This clarification is reflected in the revised text (page 5, lines 206–208):\n“However, at 7 dpi and 14 dpi, when macrophage numbers rebounded (Supplementary Figure 1m–p), FCER1G⁺ PDGFRα⁺ mesenchymal cells reappeared in the wounds (Supplementary Figure 5j–l).”\nB. Fig. 2c lacks green or red cells as indicated in the figure label.\nIn Figure 2c, the red FCER1G signal is almost entirely co-localized with the green PDGFRα signal. Thus, the merged channel image resulted in a yellow appearance due to the overlapping red and green color. To improve clarity, we have now revised Figure 2c to include split channels, allowing better visualization of each individual signal. White arrows indicate the presence of macrophages and the pink arrows indicate the presence of FCER1G⁺ PDGFRα⁺ mesenchymal cells.\nC. The Fcer1g+ cells appear to be around larger organized structures like blood vessels -- did the authors analyze whether they colocalize with CD31?\nWe performed triple immunofluorescence staining of FCER1G, PDGFRα and CD31 of the 3dpi MaFIA wound treated with vehicle, and added the following text in the Result section (page 5, lines 196-201):\n“To assess the spatial relationship between FCER1G⁺ PDGFRα⁺ mesenchymal cells and vascular structures, we performed triple immunofluorescence staining for CD31, PDGFRα, and FCER1G. FCER1G⁺ PDGFRα⁺ cells were frequently located in proximity to CD31⁺ endothelial structures but did not colocalize with them, indicating that these mesenchymal cells are perivascularly associated rather than endothelial in identity (Supplementary Figure 5h).”\n4. Depletion of Fcer1g+ cells.\nA. Since Fcer1g seems to be expressed in a subset of mesenchymal cells, why are all of the Pdgfra+ cells depleted in Fig.3b. The authors should analyze the other types of mesenchymal cells that might remain in the skin after depletion--are there any SMA+ cells?\nThe depletion strategy specifically targets FCER1G⁺ PDGFRα⁺ mesenchymal cells and does not affect PDGFRα⁺ cells that lack FCER1G expression. In the revised Figure 3b, PDGFRα⁺ single-positive cells remain detectable in 7 dpi wounds, whereas FCER1G⁺ PDGFRα⁺ double-positive cells are markedly reduced, consistent with partial depletion of the mesenchymal compartment rather than a complete loss of PDGFRα⁺ cells.\nTo evaluate additional mesenchymal subtypes, we performed immunofluorescence staining for α-smooth muscle actin (SMA) on 7 dpi wound sections. SMA⁺ cells were readily detectable in both control and FCER1G⁺ PDGFRα⁺-depleted wounds (Supplementary Figure 6d). Quantification showed that SMA⁺ cells constituted approximately 0.8% of total wound cells in both groups, with no significant difference between control and depleted wounds (Supplementary Figure 6e). These findings indicate that depletion of FCER1G⁺ PDGFRα⁺ mesenchymal cells does not broadly affect SMA⁺ mesenchymal populations.\nThis clarification has been added to the Results section (page 6, lines 232–235):\n“Immunofluorescence staining for smooth muscle actin (SMA) at 7 dpi revealed no significant difference between depleted and control wounds (Supplementary Figure 6d–e), confirming that SMA⁺ mesenchymal cells were unaffected by the depletion of FCER1G⁺ PDGFRα⁺ mesenchymal cells.”\nB. Is Fcer1g expressed in other cell types in the skin?\nWe thank the reviewer for pointing out the possibility that other cell types may also express FCER1G. Following this suggestion, we performed IHC of FCER1G on the wounded skin at 3 dpi (Supplementary Figure 5e). This labels other cell types within the skin, including macrophages, fibroblasts and leukocytes.\nC. The authors should characterize wounds of the Fcer1g depletion mouse model prior to D7-- is inflammation normal? How does epithelialization, revascularization, fibroblast repopulation occur after injury?\nWe characterized early wound healing at 3 dpi by immunofluorescence for F4/80 (macrophages), CD31 (angiogenesis), and PDGFRα (fibroblast/mesenchymal cells) in control and FCER1G⁺ PDGFRα⁺–depleted wounds (Supplementary Figure 6h–o) and added the findings to the Result section (page 6, lines 240-246):\n“At 3 dpi, immunofluorescence analysis demonstrated comparable levels of F4/80⁺ macrophages (Supplementary Figure 6h-i), CD31⁺ endothelial cells (Supplementary Figure 6j-k), and K10⁺ keratinocytes between groups (Supplementary Figure 6n-o), indicating normal inflammatory, angiogenic, and reepithelialization responses. In contrast, PDGFRα staining was reduced in depleted wounds at this early stage (Supplementary Figure 6l-m), consistent with loss of the Fcer1g⁺ mesenchymal subset.”\nThese findings further confirmed that depletion of FCER1G⁺ PDGFRα⁺ mesenchymal cells did not impair early wound healing responses, so we added additional text in the Discussion section (page 9, lines 360-372):\n“Fibroblasts comprise heterogeneous subpopulations with distinct functional roles in wound repair. For example, αSMA⁺ myofibroblasts are essential for contractile repair, as their targeted depletion completely blocks wound closure [51]. In contrast, pro-fibrotic Prrx1⁺ fibroblasts mainly drive scar formation in the ventral dermis, and their ablation reduces fibrosis without affecting closure [14]. Engrailed-1 lineage fibroblasts contribute to both repair and fibrosis; their depletion delays re-epithelialization while decreasing collagen deposition [13]. In this context, our findings show that depletion of FCER1G⁺ PDGFRα⁺ mesenchymal cells in dorsal wounds did not alter inflammatory, angiogenic, or epithelial responses during the early wound-healing phase and resulted in similar closure kinetics throughout the healing process, indicating that these cells are not required for the efficacy of wound healing. However, their loss markedly reduced scar size and collagen deposition at later stages, suggesting FCER1G⁺ PDGFRα⁺ cells as a pro-fibrotic mesenchymal subset primarily responsible for fibrosis.”\n5. I am not sure the deletion of FN1 adds much to the paper given the lack of specific depletion in macrophages and the poor characterization of the model in the paper. The immunostaining in Fig. 5 is not very convincing and the quantification is not consistent with the prior analysis.\nWe acknowledge that the current genetic model does not selectively deplete fibronectin in macrophages, which limits its specificity and may introduce discrepancies compared with the macrophage-depletion experiments. We have added additional text in the Discussion section (page 10, lines 408-411) to acknowledge this limitation:\n“However, the current EDA-fibronectin knockout model deletes fibronectin globally rather than specifically in macrophages, and thus the reduction of Fcer1g-expressing mesenchymal cells likely reflects a combined effect from loss of both macrophage- and non-macrophage-derived fibronectin.”\nWe thank the reviewer for pointing out the inconsistency with prior analyses. In addition to reporting FCER1G⁺ PDGFRα⁺ mesenchymal cell numbers per high-power field (HPF), we now include a complementary quantification normalized to the total PDGFRα⁺ cell population (Supplementary Figure 8h). This normalization was also used in our macrophage-depletion analysis (Figure 2g, Supplementary Figure 5l). At 7 dpi, macrophage-depleted MaFIA wounds contained an average of 40% FCER1G⁺ PDGFRα⁺ cells relative to 80% in vehicle-treated controls (Supplementary Figure 5l), whereas at 8 dpi, EDA-fibronectin-deficient wounds exhibited 10% compared with 35% in littermate controls (Supplementary Figure 8h). Although the absolute percentages are lower in the EDA-fibronectin model—likely due to differences in genetic background, collection time points, and the global depletion of fibronectin —the overall trend is consistent between the macrophage-depletion and EDA-fibronectin-deficient models.\nThese findings, while preliminary, suggest that fibronectin, particularly its EDA splice variant, may act upstream to promote the emergence of this pro-fibrotic mesenchymal subset.\nGiven the limited specificity of the EDA-fibronectin model and to maintain focus on the macrophage–mesenchymal interaction, we have moved the FN1 data to Supplementary Figure 8 and clarified its supportive rather than central role in the manuscript. Additionally, we strengthened the translational relevance of the study by incorporating a new human dataset in the revised Figure 5, which demonstrates conservation of FCER1G⁺ mesenchymal cells in human wound healing.\nMinor comments:\n1. It would helpful to include the annotation of the cell types from Fig. 1F in Fig. S1.\nWe agree that the labeling in the original Fig. S1 was small. To improve clarity, we have revised the figure labeling and layout in the New Supplementary Figure 4b.\nReviewer #2 (Remarks to the Author):\nThe paper explores a novel mesenchymal cell subpopulation activated by macrophages that induces fibrosis. It would be beneficial to mention whether this subpopulation exists in human skin, as the potential for this to translate into a therapy remains unclear.\nWe thank the reviewer for recognizing the novelty and potential clinical translation of our findings. In this revision, we included in silico analyses of two independent human wound-healing cohorts derived from publicly available datasets. GSE178411 includes bulkRNA-seq collected from 108 human samples comprising of uninjured skin (n=26), early wounds (n=22), late wounds (n=29), chronic wounds (n=3) and hypertrophic scars (n=28), and GSE241132 includes 12 scRNA-seq data collected from 3 human donors and sequenced at 4 different time points (day 0, 1, 7 and 30 post skin injury).\nThrough these two datasets, we demonstrated that FCER1G-expressing mesenchymal cells are also present in human wound healing, and that their temporal dynamics and transcriptional features are highly conserved between mouse and human. We added the following new section to the Results (page 8–9, lines 318–348):\n“Fcer1g-expressing mesenchymal cells are conserved in human wound healing\nTo test whether FCER1G⁺ PDGFRα⁺ mesenchymal cells are conserved in human wound healing, we reanalyzed a previously published human skin wound healing scRNA-seq dataset (GSE241132) collected at days 0, 1, 7, and 30 post-injury [50]. Within this dataset, PDGFRA and PDGFRB marked mesenchymal populations (Supplementary Figure 9a–b), which were further subclustered into 13 distinct subclusters (Figure 5a). Using gene signatures derived from mouse Fcer1g-expressing mesenchymal cells (Figure 2b), as well as FCER1G expression alone, we identified a human mesenchymal subcluster (cluster 11) that uniquely expressed both the FCER1G signature (Figure 5b-c) and FCER1G itself (Supplementary Figure 9c–d). The abundance of human FCER1G-expressing mesenchymal cells (cluster 11) peaked at day 7 post-injury (Figure 5d–e), consistent with the kinetics observed in our mouse model (Supplementary Figure 5g). FCER1G also exhibited one of the highest U-scores in human mesenchymal subcluster 11 (Supplementary Figure 9e, Supplementary Table 3). Moreover, the top differentially expressed genes (DEGs) of FCER1G⁺ PDGFRα⁺ mesenchymal cells were highly conserved between human and mouse, suggesting this represents an evolutionarily conserved mesenchymal population that responds to skin injury (Figure 5f, Supplementary Table 4).\nTo evaluate whether FCER1G⁺ PDGFRα⁺ mesenchymal cells are associated with human scar formation, we further reanalyzed another publicly available bulk RNA-seq dataset (GSE178411), which includes 108 human samples across uninjured skin (n = 26), early wounds (n = 22), late wounds (n = 29), chronic wounds (n = 3), and hypertrophic scars (n = 28). The top 100 DEGs derived from both mouse and human FCER1G⁺ PDGFRα⁺ mesenchymal cells effectively distinguished wound and scar samples from uninjured skin (Figure 5g, Supplementary Figure 9f). ssGSEA analysis revealed that FCER1G-associated signatures peaked during the early (acute) wound phase and gradually declined as wounds progressed toward hypertrophic scarring (Supplementary Figure 9g), consistent with the temporal dynamics observed in mice. Furthermore, FCER1G expression alone recapitulated this pattern (Supplementary Figure 9h), supporting its role as a key marker of a pro-fibrotic mesenchymal subset involved in hypertrophic scar formation during wound healing.”\nComments\n1. Validating FCER1G expression in human wound edge data (e.g., GSE178411) would be helpful.\nTo validate FCER1G expression in human wound tissues, we reanalyzed the publicly available bulk RNA-seq dataset GSE178411, which includes samples from uninjured skin, early and late wounds, chronic wounds, and hypertrophic scars. FCER1G expression was elevated during the early wound phase and declined over time as wounds progressed toward hypertrophic scarring (Supplementary Figure 9h). To further validate this pattern, we performed single-sample gene set enrichment analysis (ssGSEA) using FCER1G⁺ PDGFRα⁺ mesenchymal signatures derived independently from the human (GSE241132) and mouse scRNA-seq datasets. These scores were significantly enriched during the early wound phase and decreased during scar formation (Figure 5g, Supplementary Figure 9f), consistent with the kinetics observed in the mouse wounds. We added this the Result section (page 8-9, lines 336-348):\n“To evaluate whether FCER1G⁺ PDGFRα⁺ mesenchymal cells are associated with human scar formation, we further reanalyzed another publicly available bulk RNA-seq dataset (GSE178411), which includes 108 human samples across uninjured skin (n = 26), early wounds (n = 22), late wounds (n = 29), chronic wounds (n = 3), and hypertrophic scars (n = 28). The top 100 DEGs derived from both mouse and human FCER1G⁺ PDGFRα⁺ mesenchymal cells effectively distinguished wound and scar samples from uninjured skin (Figure 5g, Supplementary Figure 9f). ssGSEA analysis revealed that FCER1G-associated signatures peaked during the early (acute) wound phase and gradually declined as wounds progressed toward hypertrophic scarring (Supplementary Figure 9g), consistent with the temporal dynamics observed in mice. Furthermore, FCER1G expression alone recapitulated this pattern (Supplementary Figure 9h), supporting its role as a key marker of a pro-fibrotic mesenchymal subset involved in hypertrophic scar formation during wound healing.”\n2. The Macrophage depletion method section (Line 319-325) requires clarification. It mentions retro-orbital injection of AP20187; however, the original reference describes intraperitoneal injections. Please confirm and correct the method used for macrophage depletion.\nWe used retro-orbital venous sinus injection of AP20187 for macrophage depletion rather than the intraperitoneal route described in the original reference. This approach was chosen to avoid peritoneal adhesions, which occur in approximately 76% of MaFIA mice following intraperitoneal injection and can confound wound-healing outcomes (Burnett et al., J. Surg. Res., 2005). Retro-orbital administration is equally effective for systemic delivery and has been reported to be less stressful than intravenous injection (Steel et al., Lab Animal, 2008). To confirm the efficacy of this modified protocol, we assessed macrophage depletion at 3 days post-injury. Flow cytometry and immunofluorescence analyses demonstrated robust depletion of F4/80⁺ macrophages in the skin, as well as in the bone marrow and spleen, validating that retro-orbital AP20187 administration effectively depletes macrophages both systemically and locally.\nTo validate the efficacy of our modified protocol, we assessed macrophage depletion at 3 days post-injury. Flow cytometry and immunofluorescence analyses showed robust depletion of F4/80⁺ macrophages in the skin (Supplementary Figure 1i–l), as well as in the bone marrow and spleen (Supplementary Figure 1a–h). We added this to the Result section (page 2-3, lines 85-92) as:\n“To further confirm that GFP⁺ depletion reflects loss of macrophages, we performed flow cytometry for F4/80 and CD11b double-positive cells in the bone marrow, spleen, and skin from vehicle- and AP20187-treated MaFIA mice (Supplementary Figure 1). These analyses confirmed robust depletion of F4/80⁺CD11b⁺ macrophages in the skin (>75%) (Supplementary Figure 1i–j) and moderate depletion in the bone marrow and spleen (~50%) (Supplementary Figure 1a–b and 1e–f). Immunofluorescence staining further validated the loss of F4/80⁺ macrophages in these tissues (Supplementary Figure 1c–d, 1g–h, and 1k–l).”\nThese findings confirm that retro-orbital administration of AP20187 in MaFIA mice effectively depletes macrophages systemically and locally in the skin.\n3. The study's major limitation is its lack of validation using human data or in vitro experiments.\nWe have addressed the limitation by incorporating both human data and in vitro validation into the revised manuscript.\nAs detailed in our response to comment (1), we analyzed two independent human cohorts using orthogonal transcriptomic platforms (scRNA-seq and bulk RNA-seq) and found that the FCER1G⁺ PDGFRα⁺ mesenchymal population identified in mouse wounds is conserved in human wound healing (Figure 5 and Supplementary Figure 9). Additionally, we performed a key in vitro co-culture experiment (now shown in Figure 2i–k) to directly test whether macrophages can induce the emergence of FCER1G⁺ PDGFRα⁺ mesenchymal cells. NIH/3T3 mouse dermal fibroblasts (MDF), labeled with CellTrace™ Violet, were co-cultured with bone marrow–derived macrophages from MaFIA mice at a 1:1 ratio for 24 hours. Flow cytometric analysis revealed a distinct FCER1G⁺ PDGFRα⁺ population in the co-culture condition, which was absent in MDF cultured alone. Quantification confirmed a significant induction of this population by macrophages, supporting a direct role in driving the emergence of this mesenchymal subset. We have added this to Result section (page 5-6, lines 209-217):\n“To determine whether macrophages directly induce the emergence of FCER1G⁺ PDGFRα⁺ mesenchymal cells, we performed an in vitro co-culture assay using NIH/3T3 mouse dermal fibroblasts (MDFs) and bone marrow–derived macrophages isolated from MaFIA mice. MDFs were labeled with CellTrace™ Violet (CTV) and co-cultured with macrophages at a 1:1 ratio for 24 hours. Flow cytometric analysis of CTV⁺ MDFs revealed the appearance of a distinct FCER1G⁺ PDGFRα⁺ subpopulation in the co-culture but not in MDFs cultured alone (Figure 2i–k). Quantification confirmed a significant increase in FCER1G⁺ PDGFRα⁺ mesenchymal cells upon macrophage co-culture, indicating that macrophages directly promote the induction of this mesenchymal subset.”\n4. The Cell Culture Methods section (Lines 438–450) requires clarification. It states that mouse tails were used to isolate primary dermal mesenchymal cells. However, given the focus of the study, it would be more appropriate to isolate these cells from the wound edge tissue and perform the cell proliferation assay accordingly. If tail tissue was intentionally used, please justify this choice.\nThe reference to primary dermal mesenchymal cells isolated from mouse tails in the Cell Culture Methods section was included in error, as these cells were not used in any experiments reported in this study. We have removed this section from the revised manuscript to prevent confusion. All functional analyses were performed in vivo using wound-edge–derived cells, and for the in vitro co-culture experiment (Figure 2i–k), we used NIH/3T3 mouse dermal fibroblasts."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have addressed my prior concerns and clarified several aspects of their methods, data presentation, and conclusions. This paper is an interesting innovation in the field of regenerative biology and is a substantive advance for the field.", "reviewer_2": "The authors responded to all of our comments. Therefore, I have no issues to accept the manuscript.", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": ""}]

Decision letter