Skip to main content
NCBI home page
UKPMC Funders Author Manuscripts logoLink to UKPMC Funders Author Manuscripts
. Author manuscript; available in PMC: 2026 Jul 6.
Published in final edited form as: J Cell Biol. 2026 May 12;225(7):e202505012. doi: 10.1083/jcb.202505012

Eosinophils promote a second wave of postnatal smooth muscle differentiation in intestinal villi

Tatiana V Petrova 1,2,*, Kelly de Korodi 1, Thea Berg 1, Tania Wyss 1, Yahya Mohammadzadeh 1, Lida Safazada 1, Kathleen Shah 3, Nicola L Harris 3,+, Jeremiah Bernier-Latmani 1,*
PMCID: PMC7619177  EMSID: EMS215599  PMID: 42118023

Abstract

Intestinal villus smooth muscle cells (SMCs) play a critical structural and functional role in the gut, notably facilitating dietary fat transport via villus-associated lymphatic lacteals. In the early postnatal period subepithelial PDGFRα+ fibroblasts are guided by lacteals to differentiate and assemble into villus SMCs. We show now that villus SMC expansion can occur in the absence of lacteals suggesting the involvement of additional mechanisms. Using whole-mount imaging, genetic lineage tracing, and eosinophil-deficient models, we identify a second wave of intestinal SMC differentiation where villus tip perivascular PDGFRβ+ cells are a reservoir of distal SMC progenitors. We demonstrate that resident intestinal eosinophils act as key drivers and coordinators of TGFβ-dependent villus SMC expansion and maturation. Our findings propose a model in which PDGFRβ+ perivascular cells are progenitors for a second wave of postnatal villus SMCs and reveal a novel morphogenetic role of intestinal eosinophils as critical mediators of SMC development in the gut.

Introduction

The villus is a defining feature of the small intestine, and its finger-like structure serves to increase gut surface area for efficient nutrient absorption. These structures carry out this essential function through various specialized intestinal epithelial, stromal and immune cells which balance efficient nutrient uptake with gut barrier maintenance (Gehart and Clevers, 2019).

An example of coordinated villus cellular responses is dietary fat uptake. Dietary lipids are absorbed by enterocytes, which repackage them into large lipoprotein particles called chylomicrons (Tso et al., 2024). The chylomicrons are released basolaterally into the stroma where they enter villus lymphatic capillaries, lacteals, for systemic distribution (Bernier-Latmani et al., 2024; Bernier-Latmani and Petrova, 2017). During fat absorption villi undergo piston-like pumping thought to aid chylomicron entry into lacteals (Choe et al., 2015; Lee, 1971; Womack et al., 1987; Womack et al., 1988). This pumping is performed by a population of villus smooth muscle cells (villus SMCs, also called myofibroblasts) which are aligned parallel to the crypt/villus axis (Powell et al., 1999). Disrupted villus SMC development in mice lacking the ASE enhancer of homeodomain transcription factor Pitx2 display decreased lymphatic lipid absorption and stunted pup growth (Hu et al., 2021), highlighting the central role of villus SMCs in coordination of fat absorption.

Paracrine WNT, BMP, and Hedgehog signaling between intestinal epithelial cells and fibroblasts promotes embryonic development of villi and the muscularis mucosae enveloping the intestine (Huycke et al., 2019; Madison et al., 2005; Shyer et al., 2013; Walton et al., 2016). However, less is known about mechanisms driving development of specific villus SMC and fibroblast populations. Early postnatal villus SMC development is coordinated with lacteal lymphangiogenesis. Lacteals are formed shortly before birth when lymphatic endothelial cells sprout and migrate from submucosal lymphatic vessels into intestinal villi in response to VEGFC/VEGFR3 signaling (Kim et al., 2007; Nurmi et al., 2015; Tammela et al., 2008). Immature “star” cells, expressing the myofibroblast marker αSMA, are found at the leading edge of lacteal expansion, after which they differentiate and assemble into the “spindle”-shaped villus SMCs vertically aligned along lacteals (Hu et al., 2021). Most villus SMCs formed during the early postnatal period (before P8) are lacteal-associated and derived from subepithelial Pdgfra+ precursors. Mechanistically, lacteal/villus SMC interactions through DLL4/NOTCH3 signaling are necessary for proper patterning of this network (Sanketi et al., 2024).

However, in the adult small intestine only some villus SMCs surround lacteals, while most mature villus SMCs are not associated with lacteals (Bernier-Latmani et al., 2015; Choe et al., 2015; Hu et al., 2021). In line with this observation, most lacteal-associated villus SMC differentiation and maturation is completed by P9, while the non-lacteal associated villus SMC network continues to expand distally and horizontally from the lacteal (Hu et al., 2021; Sanketi et al., 2024). Furthermore, Pdgfra+ fibroblasts contribute little to the developing non-lacteal associated villus SMCs (Sanketi et al., 2024). Collectively, these observations suggest that distinct mechanisms drive the development of non-lacteal-associated villus SMCs.

Eosinophils have long been thought of as infection-responsive myeloid cells which repel pathogens through release of stored proteins from intracellular granules. However, the healthy intestine harbors the largest concentration of eosinophils in the body, especially in the upper small intestine (reviewed by (Gurtner et al., 2023b). Eosinophils mature upon entering the small intestine into different subpopulations which can both enhance or restrict inflammation (Arnold et al., 2018; Diny et al., 2022; Fallegger et al., 2022; Gurtner et al., 2023a; Jung et al., 2015; Kutyavin et al., 2024; Li et al., 2023; Wang et al., 2022). In mice, intestinal eosinophil density in villi increases during the postnatal period, reaching the highest density in adults (Arnold et al., 2018; Mishra et al., 1999). Adult eosinophil-deficient ΔdblGata1 mice (Yu et al., 2002) harbor smaller villi than control mice (Ignacio et al., 2022; Kutyavin et al., 2024) and, notably, eosinophil-deficient adult mice display decreased dietary lipid uptake and do not gain weight on high fat diet (Ignacio et al., 2022; Wu et al., 2011), indicating that eosinophil presence promotes dietary fat absorption.

In this work, we show that Pdgfrb+ perivascular cells are an alternate source for villus SMCs and that eosinophils are necessary for a second wave of villus SMC expansion, differentiation and maturation in the postnatal small intestine. As eosinophils migrate into villi, they actively contribute to the growing villus SMC network and myogenic program. Eosinophils express Tgfb2 and eosinophil deficiency significantly decreases direct TGFβ target gene expression in pup villus fibroblasts. As TGFβ signaling blockade and specific Tgfbr2 ablation in Pdgfrb+ fibroblasts in pups phenocopies villus SMC defects in the absence of eosinophils we propose that eosinophil-derived TGFβ is crucial for later postnatal villus SMC expansion and maturation.

Results and Discussion

A second wave of postnatal villus SMC expansion in the later postnatal period

Sanketi et al. recently reported that by P9 most villus SMCs were lineage-traced from subepithelial Pdgfra+ fibroblasts when labeled from P0, but there were significantly fewer traced villus SMCs when labeling was started at P9. Furthermore, the P0 Pdgfra lineage-traced SMCs mostly accumulated around lacteals, but not among non-lacteal associated villus SMCs (Sanketi et al., 2024). These results indicate that specific villus SMC subset differentiation may be controlled in a temporal manner from distinct cellular sources to inhabit discrete villus zones. Hu et al. reported αSMA+ star-shaped SMC precursor cells near villus blood capillaries (Hu et al., 2021), and we confirmed this observation (Fig. 1A). Based on this spatial proximity, we hypothesized that perivascular mural cells could also serve as alternative precursors for villus SMCs.

Figure 1. A second wave of postnatal villus SMC expansion in the later postnatal period.

Figure 1

Open in a new tab

(A) αSMA+ star cells (arrowheads) at the villus tip are located near villus blood capillaries. Staining for αSMA+ villus SMCs (cyan) and blood capillaries (red, VEGFR2) in a P7 pup wild-type Balb/c villus. (B) Pdgfrb+ cells broadly cover villus capillaries in adult mice. Staining for Pdgfrb-GFP+ perivascular cells (green) covering all villus vessels (red, VEGFR2) including villus venules (blue, MAdCAM1) in adult mice. (C) Early postnatal villus blood capillaries are less covered by Pdgfrb+ cells than adults. Staining for mGFP (green) expressed by Pdgfrb+ cells covering villus capillaries (red, PECAM1) 48h after activation by injection of 4OHT. (D) Scheme for lineage tracing experiment to test Pdgfrb+ cell incorporation into the developing villus SMC network using littermate Pdgfrb-CreERT2; mTmG pups for each timepoint. (E-G) Perivascular Pdgfrb+ cells are incorporated into the growing villus SMC network postnatally. (E) Staining for the developing villus SMC network (red, αSMA; blue, CNN1) and Pdgfrb+ cells (green, GFP) during lineage tracing for 24h (P6), 48h (P7) and 5 days (P10) in Pdgfrb-CreERT2; mTmG pups. Arrowheads, GFP+ villus SMCs. (F-G) Quantification of the (F) number of GFP+ αSMA+ SMCs per villus area and (G) percent of GFP+ villus SMCs present in the villus bottom, middle or top as shown on the scheme, n=7 pups, P6; n=4 pups, P7 and n=7 pups, P10. (H) Scheme for postnatal control and anti-VEGFR3 antibody treatment in C57Bl/6 pups. (I) Postnatal villus SMCs are present in the absence of lacteals. Staining for villus SMCs (αSMA, red on top, white on bottom) and lacteals (green, LYVE1) in P10 pups treated either with control or anti-VEGFR3 blocking antibodies. Arrowheads, star αSMA+ cells. Scale bars: 50μm: B, C, E, I; 20μm: A. All values shown as mean ± SD. **P<0.01, ****P<0.0001, 1-way ANOVA with Tukey’s post-hoc test for multiple comparisons.

The majority of perivascular cells are PDGFRβ+ (Muhl et al., 2020) and PDGFRβ+ cells are associated with villus capillaries in adult mice and pups (Hong et al., 2020)(Fig. 1B, C), a subset of which are likely pericytes. We therefore performed lineage tracing in Pdgfrb-CreERT2; mTmG pups (Cuervo et al., 2017; Muzumdar et al., 2007) by injecting 4-OHT at P5 and analyzing at P6, 7 and 10 to determine whether αSMA+GFP+ cells incorporated into the villus spindle SMC network (Fig. 1D). After 24h of lineage tracing most lineage-traced cells were perivascular, while few were detected in subepithelial zones (Fig. 1E; Fig. S1A). While perivascular GFP+ cells were readily observed at P6 and P7, very few αSMA+GFP+ spindle SMCs were detected (Fig. 1E, F). However, by P10 the number of αSMA+GFP+ spindle cells was significantly increased (Fig. 1E, F) indicating that PDGFRβ+ cells contribute to the maturing spindle villus SMC network at later stages of postnatal development. Lineage-traced GFP+ cells were distributed throughout the villus, however spindle-shaped GFP+ villus SMCs were significantly enriched at the villus top rather than the bottom (Fig. 1E-G), demonstrating the preferential incorporation of PDGFRβ+ progenitor-derived cells into non-lacteal-associated SMCs at the villus tip.

Lacteal to villus SMC Notch signaling is necessary for early postnatal villus SMC expansion (Sanketi et al., 2024). To test if this mechanism was required in the later postnatal period we inhibited lacteal expansion by treating pups with blocking antibodies against VEGFR3 (mF4-31C1)(Pytowski et al., 2005) from P1 to P10 (Fig. 1H). Despite the absence of lacteals, P10 mice treated with VEGFR3 antibody displayed spindle-shaped villus SMCs similar to those in pups treated with control IgG. (Fig. 1I). Moreover, villus tip αSMA+ star precursor cells were observed in pups without lacteals (Fig. 1I), validating our hypothesis for an alternative mechanism for expansion of the postnatal villus SMC network at later stages of development.

Taken together, these data suggest a model where, along with subepithelial Pdgfra+ fibroblasts (Sanketi et al., 2024), perivascular Pdgfrb+ cells also can become mature villus SMCs, albeit in a temporally and spatially distinct manner. Moreover, we show that αSMA+ star precursor cells persist in the absence of lacteals suggesting the existence of alternative drivers of villus SMC maturation and expansion. This developmental redundancy may underlie functional importance to properly develop villus SMCs to promote efficient nutrient absorption and structural integrity.

Eosinophil postnatal recruitment pattern and cellular interactome suggest a role in villus SMC development

Adult eosinophil-deficient ΔdblGata1 mice exhibit smaller villi than control mice (Ignacio et al., 2022; Kutyavin et al., 2024), suggesting that intestinal eosinophils play a role in villus stroma development. Analysis of the ECM proteins fibronectin and tenascin C, which are broadly expressed in villus stromal cells (Bernier-Latmani et al., 2015; Bernier-Latmani et al., 2022; Bernier-Latmani and Petrova, 2016), showed that the main villus defect of eosinophil deficiency is loss of stroma size (Fig. 2A). Interestingly, the reduced villus size in ΔdblGata1 mice becomes apparent only by P10 (Ignacio et al., 2022), coinciding with the second wave of villus SMC expansion.

Figure 2. Eosinophil postnatal recruitment pattern and cellular interactome suggest a role in villus SMC development.

Figure 2

Open in a new tab

(A) Villus stroma is smaller in the absence of eosinophils. Staining for fibronectin (red, top) and tenascin C (green, bottom) in intestinal villi from control Balb/c and eosinophil-deficient ΔdblGata1 adult mice. (B) Progressive migration of eosinophils into the postnatal villus. Staining for blood vessels (red, VEGFR2) and eosinophils (green, SIGLECF) in intestinal villi at several postnatal timepoints. Quantification of distance of individual eosinophils from the villus base (as a % of villus height) at each timepoint. Each dot is one eosinophil measurement; P0.5 represents data from 2 individual pups, P4, P7, P10 represent data from 3 individual pups, 10 weeks represents data from 4 individual mice. (C-D) Quantification of (C) villus eosinophil density and (D) eosinophil height along the villus at several postnatal timepoints, n=2, P0.5; n=3, P4, 7 and 10; n=4, 10 weeks. (E) Serial sections from a scanning block-face electron microscopy (SBEM) image stack from an adult wild-type mouse villus. Colors hand-drawn onto the image based on cell type: green, eosinophils; cyan, smooth muscle cells; yellow, fibroblasts; magenta, phagocytic cells; violet, enteric glial cells; red, blood vessel; arrows, eosinophil; asterisks, direct eosinophil contact with other cells. (F) Quantification of SBEM images for direct contacts of eosinophils with other stromal cells, n=3 mice. (G) Eosinophils are in contact with Pdgfrb lineage-traced GFP+ cells. Staining for GFP (green), αSMA (red) and eosinophils (white, SIGLECF) in Pdgfrb-CreERT2; mTmG pups lineage traced from P5 to P7. (H) Timeline for villus SMC expansion in the postnatal small intestine. From the late embryonic period to P7 villus SMCs are lacteal-associated and derived from Pdgfra+ villus fibroblasts. Our model proposes that after P7 non-lacteal associated villus SMCs are expanded from Pdgfrb+ perivascular cells, a process promoted by the presence of eosinophils. Scale bars: 50μm: A, B; 20μm: G; 5μm: E. All values shown as mean ± SD.

To gain insight into temporal and spatial patterning of eosinophil entry into intestinal villi we analyzed the abundance and location of villus eosinophils postnatally. As expected (Ignacio et al., 2022; Mishra et al., 1999), few gut eosinophils were detected in E17.5 control embryos, while no SIGLECF-high eosinophils were detected in ΔdblGata1 embryos (Fig. S1B). Scarce villus intestinal eosinophils were detected at P0.5, however, the few present were restricted to the proximal villus base. By P4 and P7 eosinophil numbers increased, but the majority were still restricted to the villus base (Fig. 2B). In contrast, at P10, eosinophils were more evenly distributed along the villus axis, approaching abundance and distribution of adult 10-week-old mice (Fig. 2B-D). Therefore, eosinophils enter the villus from the base and progress from just after birth to completely fill the villus to the tip by P10.

To interrogate cells in proximity to intestinal eosinophils we performed serial-blockface scanning electron microscopy on villi from adult wild-type mice allowing precise 3D analysis of cell-cell interactions. Eosinophils were identified as cells displaying abundant cytoplasmic granules and harboring distinctive lobular nuclei (Spencer et al., 2014) and had most direct cell-cell contacts with fibroblasts, followed by other eosinophils and phagocytic cells, likely macrophages, and importantly villus SMCs (Fig. 2E, F; Video 1). In addition, eosinophils were observed in contact with Pdgfrb lineage-traced GFP+ star cells at P7 (Fig. 2G), suggesting they play an active role in later villus SMC expansion. Since the P7-P10 timeframe corresponded to non-lacteal associated villus SMC expansion and development we hypothesized that eosinophils contribute to this process (Fig. 2H).

Eosinophils promote postnatal villus smooth muscle cell expansion and differentiation

Since eosinophils are in contact with villus SMCs (Fig. 2E, F), we hypothesized that eosinophils play a direct role in villus SMC development. Indeed, confocal microscopy confirmed a close association between eosinophils and villus SMCs in adult wild-type mice (Fig. 3A, S1C). In addition, adult ΔdblGata1 mice displayed fewer villus SMCs than controls (Fig. 3A, B)(Ignacio et al., 2022) and decreased villus SMC abundance was restricted to non-lacteal associated villus SMCs (Fig. 3C). Immunostaining for αSMA in adult and P10 pups revealed a significantly reduced network of villus SMCs in ΔdblGata1 pups (Fig. 3D) indicating that eosinophils contribute to postnatal villus SMC network expansion.

Figure 3. Eosinophils promote postnatal villus smooth muscle cell expansion and differentiation.

Figure 3

Open in a new tab

(A) Eosinophils are associated with villus SMCs in adult mice. Staining for eosinophils (green, SIGLECF) and villus SMCs (cyan, αSMA) in Balb/c and ΔdblGata1 adult mice. (B) Reduced density of villus SMCs in adult eosinophil-deficient adult mice. Staining for eosinophils (green, SIGLECF), villus SMCs (cyan, αSMA) and blood capillaries (red, VEGFR2) in Balb/c and ΔdblGata1 adult mice. (C) Eosinophil deficiency leads to loss of non-lacteal associated villus SMCs. Staining for villus SMCs (cyan, αSMA) and lacteals (green, LYVE1; red CCL21) in Balb/c and ΔdblGata1 adult mice. (D) Decreased villus SMC network in eosinophil-deficient pups. Staining for eosinophils (green, SIGLECF) and villus SMCs (cyan, αSMA) in littermate Balb/c and ΔdblGata1 P10 pups. Quantification of villus SMC area per villus area in Balb/c and ΔdblGata1 P10 pups, n=6 for each genotype, combined data from two independent experiments. (E, F) Villus SMC maturation corresponds with eosinophil entry into the postnatal villus. Staining for eosinophils (green, SIGLECF), villus SMCs (cyan, αSMA) and blood capillaries (red, VEGFR2) in Balb/c pups at (E) P2 and (F) P4 and P10. Inset highlights interaction between eosinophils and star αSMA+ cells. (G) αSMA+ star cells mature through a αSMA+ CNN1low “transition” state before becoming αSMA+ CNN1+ spindle cells. Staining for αSMA (red) and CNN1 (green) showing villus tip star cells (arrows), transition cells (arrowheads) and spindle cells (αSMA+CNN1+) in P7 Balb/c pups. (H-J) Decreased number of transition cells in the absence of eosinophils. (H) Staining for αSMA (red) and CNN1 (green) in littermate Balb/c and ΔdblGata1 P7 pups, star cells are highlighted with arrowheads. (I-J) Quantification of (I) transition and (J) star cells in Balb/c and ΔdblGata1 P7 pups, n=5 for each genotype, combined data from two independent experiments. Scale bars: 50μm: B, C, D, E, F, H (whole villus); 20μm: A, G, H (villus tips). All values shown as mean ± SD. **P<0.01, ***P<0.001, 2-tailed unpaired Student’s t test.

The αSMA+ star cells present in villi right after birth mature into elongated spindle cells extending from the villus base (Hu et al., 2021). Since eosinophils also enter the villus base postnatally (Fig. 2B)(Ignacio et al., 2022; Mishra et al., 1999), we hypothesized that villus SMC maturation and eosinophil migration were cotemporaneous. Analysis of postnatal villus SMCs and eosinophils showed coordinated development of the SMC network and eosinophil migration to the distal villus tips, with a full spindle shape network extending to the villus tips by P10, coincident with the presence of eosinophils (Fig. 3E, F).

αSMA+ spindle cells express contractile proteins found in mature muscle cells, while star cells do not (Hu et al., 2021). We therefore analyzed postnatal villus SMCs using calponin-1 (CNN1) which is expressed in mature, contractile SMCs (Wang et al., 2015). Analysis of P7 villus tips for αSMA and CNN1 showed that most star cells were αSMA+CNN1- and the main spindle cell population displayed high expression of both αSMA and CNN1. However, we observed low CNN1 expression in a proportion of star cells lower in the villus, near the spindle cell population, which we term “transition” cells (Fig. 3G). At P7, transition αSMA+CNN1low cells were readily observed in villi of control pups but were significantly reduced in ΔdblGata1 pups (Fig. 3H, I), while the number of αSMA+CNN1- star cells was not changed (Fig. 3H, J). These data indicate that eosinophils facilitate villus SMC expansion by promoting star-like precursor cell differentiation towards a spindle villus SMC fate.

In all, these results show that early postnatal eosinophil colonization is necessary for villus SMC expansion and differentiation. Eosinophils have long been associated with increased fibrosis and stromal cell remodeling in pathological states including eosinophilic esophagitis and asthma (Furuta and Katzka, 2015; Papi et al., 2018). In the adult gut eosinophils increase pathological expansion of submucosal SMCs during radiation-induced fibrosis (Takemura et al., 2018). However, here we show that eosinophils also promote SMC formation as part of a controlled, developmental program. Interestingly, in the postnatal lung there is a surge of eosinophil infiltration which promotes extracellular matrix remodeling, including enhancing tenascin C deposition (Loffredo et al., 2020). In addition during mammary gland development eosinophils promote duct branching (Gouon-Evans et al., 2000). Therefore, eosinophils may be more active as postnatal stromal organizers than previously recognized.

Interestingly, disruption of villus SMC development in Pitx2ASE/ASE mice was associated with decreased fat uptake in pups (Hu et al., 2021). Consistent with those observations, eosinophil-deficient adult mice also display reduced fat absorption (Ignacio et al., 2022; Wu et al., 2011). Therefore, eosinophil-mediated SMC patterning is likely functionally important for adults nutrient absorption, perhaps through promoting piston-like villus movements that propel gut lymphatic flow (Bernier-Latmani and Petrova, 2017).

Intestinal eosinophils preferentially express Tgfb2 and promote a myogenic gene expression program in gut fibroblasts

We next sought to identify how intestinal eosinophils promote villus SMC maturation. The BCR co-receptor CD22 is mainly expressed in B cells, but is also selectively expressed in intestinal eosinophils (Kutyavin et al., 2024; Li et al., 2023; Wen et al., 2012). Notably, in adult mice its expression is increased in villus eosinophils closer to the villus tip downstream of retinoic acid signaling (Kutyavin et al., 2024). Since villus SMC differentiation correlated with eosinophil migration towards the villus tip (Fig. 3E-J), we hypothesized that CD22+ eosinophils were present at the site of SMC differentiation. SIGLECF/CD22 co-staining in P4 pups showed that CD22+ eosinophils were found throughout the villus and crypt area (Fig. 4A). Moreover, CD22+ eosinophils were associated with the “leading edge” of expanding star villus SMCs (Fig. 4A) suggesting these specific gut eosinophils play a role in villus SMC expansion and/or differentiation.

Figure 4. Intestinal eosinophils preferentially express Tgfb2 and promote a myogenic gene expression program in gut fibroblasts.

Figure 4

Open in a new tab

(A) CD22+ eosinophils are at the leading edge of villus SMC network expansion. Staining for CD22 (green), SIGLECF (blue) and αSMA (cyan) in P4 villi of wild-type C57Bl/6 pups. Visualization of CD22+ eosinophils (red) was done by masking CD22 by SIGLECF. Arrowheads, CD22+ eosinophils on the villus SMC leading edge highlighted by star cells. (B, C) Gut resident CD22+ eosinophils express Tgfb2. Volcano plots showing differential gene expression between either (B) lung and gut eosinophils (Wen et al., 2012) or (C) CD22- and CD22+ gut eosinophils (Kutyavin et al., 2024). (D) Distinct gene expression changes in intestinal fibroblasts of eosinophil-deficient pups. Unsupervised hierarchical clustering of genes differentially expressed between intestinal fibroblasts from P9 littermate Balb/c and ΔdblGata1 pups, n=3 of each genotype. (E) Expression of TGFβ target genes are reduced in intestinal fibroblasts from eosinophil-deficient pups. Volcano plot of significantly differentially expressed genes between fibroblasts sorted from P9 Balb/c and ΔdblGata1 pups. TGFβ target genes highlighted in green. (F) Normalized enrichment scores (NES) obtained through gene set enrichment analysis shows gene expression promoting myogenesis is decreased in pup fibroblasts from eosinophil-deficient mice. “Hallmark” pathways significantly differentially enriched in intestinal fibroblasts from P9 Balb/c and ΔdblGata1 pups. Scale bars: 50μm.

To assess mechanistically how gut eosinophils promote villus SMC expansion and differentiation we re-analyzed two independent RNAseq datasets from sorted intestinal eosinophils (Kutyavin et al., 2024; Wen et al., 2012) for genes coding secretory signaling proteins expressed in CD22+ gut eosinophils. We found that in the two bulk RNAseq datasets, total gut and CD22+ gut eosinophils expressed similar genes including Cd22, Clec4a4, Ahrr, Jaml, Cyp1b1 and Cdh17, compared to paired lung and integrin α4β7-CD22- eosinophils which expressed similar markers including Sell, Clec4e, Ccl9, Tnf and Il6 (Fig. 4B, C; Table S1, S2). Of note, among genes encoding secretory proteins, Tgfb2 was the most highly expressed in CD22+ gut eosinophils and it was also significantly enriched in total gut eosinophils compared to those in the lung (Fig. 4B, C). Eosinophils are known to harbor TGFβ in granules (Gigon et al., 2023) suggesting that CD22+ gut eosinophils are sources for TGFβ ligands.

To investigate how villus fibroblasts are affected by the absence of eosinophils we performed bulk RNAseq on P9 intestinal CD45-EPCAM-CD31-PDPN+ fibroblasts from control and ΔdblGata1 littermates (Fig. S1D). Analysis revealed 329 genes differentially expressed between gut fibroblasts from eosinophil-proficient and -deficient pups (Fig. 4D, E; Table S3). Of note, smooth muscle-specific genes, including Tagln, Mylk, Cnn1, Acta2, Actg2 and Myh11 were all among the most significantly downregulated in ΔdblGata1pups (Fig. 4D, E; Table S3). These genes are all direct TGFβ target genes identified in both in vivo and in vitro screens (McBryan et al., 2007; Plasari et al., 2009), showing that TGFβ signaling is reduced in the absence of villus eosinophils. Gene set enrichment analysis (GSEA) showed that several pro-inflammatory programs were increased in ΔdblGata1 pups (Fig. 4F; Table S4), consistent with loss of TGFβ signaling (Massagué and Sheppard, 2023) and increased inflammation observed in adult eosinophil-deficient mice (Gurtner et al., 2023a; Ignacio et al., 2022). Furthermore, among the pathways downregulated were that of myogenesis and oxidative phosphorylation (Fig. 4F; Table S4), hallmarks of smooth muscle cells (Hargreaves and Spriet, 2020), consistent with our observations of decreased villus SMCs in eosinophil-deficient pups. Therefore, these data suggest that intestinal eosinophils promote postnatal villus SMC expansion and maturation through enhancing TGFβ signaling.

PDGFRβ+ fibroblast-specific TGFβ signaling promotes villus SMC and villus size expansion

TGFβ signaling is the main pathway driving fibroblast to myofibroblast differentiation and maturation (Frangogiannis, 2020). Therefore, we hypothesized that eosinophil-derived TGFβ signaling drives villus SMC formation in postnatal villi. We first tested if TGFβ promotes villus SMC expansion and administered control and TGFβ blocking antibodies (1D11.16.8, binds TGFβ1, 2 and 3) to P4 wild-type pups and harvested the gut at P10 (Fig. 5A). Control antibody-treated mice displayed a villus SMC network mostly consisting of spindle cells positive for αSMA and CNN1, with scattered star and transition cells near the villus tip (Fig. 5B). In contrast, anti-TGFβ treated pups displayed significantly fewer αSMA+CNN1low transition cells and αSMA+CNN1+ cells were found significantly lower in the villus suggesting impaired villus SMC maturation (Fig. 5B-D). However, TGFβ signaling blockade did not reduce the number of villus eosinophils (Fig. 5B, E). These results show that TGFβ signaling enhances the abundance and maturation of villus SMCs in the postnatal gut.

Figure 5. TGFβ and intestinal eosinophils promote conversion of Pdgfrb+ fibroblasts to villus SMCs.

Figure 5

Open in a new tab

(A) Experimental outline of pup treatment with control or TGFβ blocking antibodies. (B-E) TGFβ signaling blockade inhibits postnatal villus SMC expansion and maturation. (B) Staining for αSMA (red), CNN1 (green) and SIGLECF (white) in villi of P10 C57Bl/6 pups treated either with control or TGFβ blocking antibodies. Arrowheads, star cells; dots, villus outline. (C-E) Quantification of (C) the number of transition cells per villus, (D) height of CNN1+ villus SMCs relative to the villus height and (E) the number of eosinophils per villus area, n=6 control IgG treated pups and n=7 TGFβ blocking antibody treated littermate pups, data combined from two independent experiments. (F) Experimental layout for postnatal ablation of TGFBR2 in Pdgfrb+ intestinal fibroblasts from P5 to P10. (G) Whole-mount immunostaining of P10 intestine for villus SMCs (red, aSMA and green, CNN1) and blood capillaries (blue, VEGFR2) from Tgfbr2fl/fl and Tgfbr2fl/fl; Pdgfrb-CreERT2 pups. White boxes denote magnified villus tips in right panels. (H-K) Quantification of (H) CNN1+ villus SMC height as a percentage of villus height; (I) number of transition aSMA+ cells per villus; (J) the aSMA+ area per villus; (K) villus area. n=4 Tgfbr2fl/fl and n=5 Tgfbr2fl/fl; Pdgfrb-CreERT2 pups. Data are representative of 2 independent experiments using littermate pups. (L) Experimental layout for ex vivo co-culture experiment with Pdgfrb-GFP intestinal fibroblasts and gut eosinophils. (M) Immunostaining of Pdgfrb-GFP intestinal fibroblasts (green, GFP) alone and co-cultured with gut eosinophils, TGFβ1 or TGFβ2 with or without the addition of SB431542; red, aSMA; blue, DAPI. (N) Quantification of the percentage area αSMA+ among conditions of Pdgfrb-GFP intestinal fibroblasts. Data combined from two independent experiments, n=3 for each treatment. Scale bars: 100μm, M; 50μm, B, G; 20μm, B, G insets. All values shown as mean ± SD. *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001 2-tailed unpaired Student’s t test.

Lineage tracing with Pdgfrb-CreERT2; mTmG pups showed that after P5 Pdgfrb+ perivascular fibroblasts contribute to the upper developing villus SMC network (Fig. 1E-G). Therefore, we tested whether TGFβ signaling in Pdgfrb+ fibroblasts is important for the developing villus SMC network. We generated Tgfbr2fl/fl; Pdgfrb-CreERT2 mice, induced Tgfbr2 deletion at P5 and harvested the pup intestines at P10 (Fig. 5F). Like TGFβ blocking antibody treatment, Tgfbr2fl/fl; Pdgfrb-CreERT2 pups displayed significantly fewer αSMA+CNN1low transition cells and mature αSMA+CNN1+ cells were found significantly lower in the villus compared to littermate controls (Fig. 5G-I). Moreover, similar to our observations in eosinophil-deficient ΔdblGata1 pups (Fig. 3D)(Ignacio et al., 2022), the villus SMC network and villus size were significantly smaller in Tgfbr2fl/fl; Pdgfrb-CreERT2 pups (Fig. 5G, J, K). These results show that Pdgfrb+ perivascular fibroblasts depend on TGFβ signaling for both villus SMC network development and villus size expansion during the later postnatal period.

Eosinophil-derived TGFβ promotes conversion of intestinal PDGFRβ+ fibroblasts to SMCs

To determine if intestinal eosinophils directly convert Pdgfrb+ gut fibroblasts to an SMC phenotype we performed ex vivo co-culture experiments. We first plated sorted GFP+ intestinal fibroblasts from Pdgfrb-GFP mice (Fig. 5L, S1E)(Gong et al., 2003). After 3 days the adherent GFP+ cells displayed a stellate shape and low to absent αSMA expression (Fig. 5M). On day 3 we added intestinal eosinophils (Fig. S1F) to Transwells in the wells containing Pdgfrb-GFP intestinal fibroblasts (Fig. 5L). To additional wells we also added TGFβ1 and TGFβ2 as positive controls. After 3 days of co-culture with intestinal eosinophils, Pdgfrb-GFP+ fibroblasts lost their stellate shape and significantly increased αSMA expression to levels equivalent with that of TGFβ1- and TGFβ2-treated fibroblasts (Fig. 5M, N). Importantly, eosinophil-mediated phenotypic changes to Pdgfrb-GFP+ were TGFβ-dependent as addition of the small molecule inhibitor of TGFβ signaling, SB431542, completely blocked αSMA expression (Fig. 5M, N).

To assess if there was decreased fat absorption after loss of TGFβ signaling we measured weights of P10 pups treated with TGFβ blocking antibodies or Tgfbr2fl/fl; Pdgfrb-CreERT2 mice. Neither treatment resulted in weight loss compared to littermate controls (Fig. S2A, B), suggesting villus SMC reduction is not critical for postnatal fat absorption. Dietary fat is absorbed in the form of chylomicrons through lymphatic vessels in intestinal villi, lacteals, and the presence of lacteals in adult mice promotes fat absorption (Bernier-Latmani et al., 2015; Nurmi et al., 2015; Roci et al., 2025; Suh et al., 2019; Zhang et al., 2018). Therefore, we tested if lacteals are necessary for postnatal fat absorption. We inhibited lacteal growth into villi by injecting VEGFR3 blocking antibodies postnatally (Fig. S2C, D) and tracked pup weight. Surprisingly, in the absence of lacteals pup weight gain was not affected (Fig. S2E).

We therefore sought to investigate an alternate functional consequence of decreased villus SMC numbers after TGFβ blockade. Interestingly, macrophages, T cells and FOXP3+ Tregs are all decorating villus SMCs in adult animals (Bernier-Latmani et al., 2015; Bernier-Latmani and Petrova, 2016)(Fig. S2F-H), suggesting that the non-lacteal associated SMC network acts as a scaffold for immune cell interactions. Therefore, we hypothesized that decreased villus SMC numbers could affect the distribution of villus macrophages postnatally. However, we observed no difference in their density, nor morphology at the villus tip (Fig. S2I), suggesting villus SMCs do not alter villus macrophage patterning or phenotype in the postnatal period.

This work shows that in contrast to the 1st wave of villus SMC expansion derived from subepithelial Pdgfra+ fibroblasts, eosinophils promote a second wave of villus SMC expansion from perivascular Pdgfrb+ fibroblasts (Fig. S3). This developmental redundancy may underlie the functional importance to properly develop villus SMCs to promote efficient nutrient absorption and structural integrity after birth. Mechanistically, we show that intestinal TGFβ, likely eosinophil-derived, drives conversion of Pdgfrb+ fibroblasts to a villus SMC phenotype and also likely contributes to other pup villus fibroblasts as villus size was significantly smaller in Tgfbr2fl/fl; Pdgfrb-CreERT2 pups. Therefore, these new insights show that intestinal eosinophils drive a developmental stromal remodeling program necessary for proper villus development.

Our work reveals that intestinal villus SMCs utilize diverse mesenchymal sources for their postnatal development and assembly. Lacteal-associated SMCs arise early during development and appear to require lymphatic endothelial-derived signals for recruitment and differentiation of PDGFRα+ subepithelial fibroblasts (Sanketi et al., 2024). We show that further expansion of villus SMCs derives from PDGFRβ+ perivascular cells, whose TGFβ-dependent differentiation is orchestrated by the intestinal eosinophils that colonize intestinal villi postnatally. These findings are consistent with previous work showing that adult mice with eosinophils also maintain larger villi than eosinophil-deficient mice (Ignacio et al., 2022; Kutyavin et al., 2024), suggesting the villus SMC network reinforces villus robustness as well as promoting villus contraction for efficient nutrient absorption.

Although eosinophils have long been known to promote TGFβ signaling through release of the ligand from secretory granules, it is most often thought to be associated with upper airway and digestive tract inflammation associated with asthma and eosinophilic esophagitis (Aceves et al., 2007; Kariyawasam and Robinson, 2007). In contrast, here we show that during the early postnatal period, both eosinophils and TGFβ signaling promote specific and controlled patterning of villus SMCs. It remains to be determined exactly how developmental eosinophil-mediated TGFβ signaling differs from pathological eosinophilia leading to tissue fibrosis. Eosinophils secrete granules by various mechanisms (Fettrelet et al., 2021) but how degranulation differs in adult inflammatory disorders vs developmental settings remains to be examined. Moreover, although CD22+ eosinophils highly express Tgfb2, release of pre-stored granule-packaged TGFβ1 could also contribute to postnatal villus SMC maturation. Indeed, mice with eosinophil-specific Tgfb1 ablation displayed increased gut inflammation (Fallegger et al., 2022), similar to observed higher inflammation in gut fibroblasts in the absence of eosinophils (Fig. 4F). However, it remains to be determined if TGFβ blockade or ablation of Tgfb1 or Tgfb2 from gut eosinophils impacts postnatal villus SMC expansion and maturation or fat uptake disruption in adults.

Latent TGFβ is mainly activated through integrin-mediated “pulling” of inhibitory latency-associated peptides (LAPs) from TGFβ rendering the peptide available for receptor binding (Massagué and Sheppard, 2023). TGFβ1+3 are activated by receiver cells expressing integrin αVβ6 or αVβ8, while TGFβ2 is only activated by receiver cell integrin αVβ6 when anchored to the donor cell by milieu anchoring proteins (like GARP(LRRC32) or LRRC33)(Le et al., 2023). It is interesting to speculate that the complex interactions necessary for specific TGFβ isoform activation may make discrete pockets where either ECM- or cell-bound TGFβ2 signaling could occur. Recent work showed that selective integrin αVβ8 expression by brain glial cells and neurons spatially restricted TGFβ-mediated fibroblast to myofibroblast conversion in the brain (Ewing-Crystal et al., 2025). Therefore, further work interrogating cell type-specific expression of TGFβ activating enzymes and potential expression of milieu anchoring proteins on eosinophils, as well as eosinophil-specific ablation of Tgfb2, could expand the findings of the current study.

We examined if TGFβ inhibition or a lack of lacteals inhibited postnatal fat absorption but found no defects in either situation. Two explanations may account for the lack of an observable postnatal phenotype. First, fully developed lacteals and villus SMCs may be particularly important under physiologically challenging conditions, such as when maternal nutrition is limited, or in the case of large litter size. In our experiments, litter sizes were small and milk production was likely sufficient, reducing the need for maximal energy extraction from absorbed milk and therefore masking potential weight differences. Second, a complete villus SMC network may become critical during the transition to solid food, which has lower nutritional density than milk. Consistent with this idea, adult eosinophil-deficient mice display decreased dietary lipid uptake (Ignacio et al., 2022; Wu et al., 2011). Thus, the postnatal expansion of the eosinophil-dependent villus SMC network may represent a developmental preparation for adult nutrient absorption.

Apart from nutrient absorption another main function of intestinal villi is immunosurveillance of the gut barrier. Macrophages, T cells and FOXP3+ Tregs all are in close proximity to villus SMCs (Bernier-Latmani et al., 2015; Bernier-Latmani and Petrova, 2016) and we hypothesize that the villus SMCs produce extracellular matrix proteins that favor immune cell attachment and could be used for facilitating cell-cell interactions. However, we also observed no postnatal phenotype in macrophage patterning in mice with TGFβ blockade suggesting that postnatal villus SMC development could be allowing immune cell distribution in preparation for solid food. Therefore, in contrast to direct secretion of immunosuppressive factors, eosinophil-mediated patterning of the villus SMC network could suppress gut inflammation or modulate immune responses by providing a platform for immunosurveillance of normal gut microbiota. Follow-up studies should include an indexing of fibroblast and immune cellular composition in eosinophil-deficient and/or TGFβ blockade along with experiments to determine postnatal and adult physiological defects.

Methods

Animal models

Animal experiments were approved by the Animal Ethics Committee of Vaud, Switzerland. ΔdblGata1 (RRID:IMSR_APB:1521), Pdgfrb-GFP (RRID:MMRRC_031796-UCD), Pdgfrb-CreERT2 (RRID:IMSR_JAX:030201), Tgfbr2fl/fl (RRID:IMSR_JAX:012603), mTmG (RRID:IMSR_JAX:007676) mice were described (Cuervo et al., 2017; Gong et al., 2003; Levéen et al., 2002; Muzumdar et al., 2007; Yu et al., 2002). ΔdblGata1 mice were maintained on a Balb/c background and Pdgfrb-GFP, Pdgfrb-CreERT2; mTmG and Tgfbr2fl/fl; Pdgfrb-CreERT2 mice were maintained on a C57Bl/6 background. C57Bl/6 and Balb/c mice were purchased from Janvier Labs. Adult mice were 8–12-weeks-old at sacrifice, pup ages were as described in the text. Pdgfrb-CreERT2; mTmG mice were injected with 4-OHT (Tocris, 50mg/kg mouse) dissolved in sunflower oil (Sigma). For VEGFR3 blockade experiments mice were injected with either anti-VEGFR3 (mF4-31C1, Eli Lilly) or control rat IgG2a (BioXcell) antibodies (40mg/kg mouse). For TGFβ blockade experiments pups were injected with TGFβ blocking (1D11.16.8) or control (IgG1) antibodies (20mg/kg mouse), both from BioXcell. Tgfbr2fl/fl and Tgfbr2fl/fl; Pdgfrb-CreERT2 pups were injected with tamoxifen diluted in Kolliphor (Sigma, 50mg/kg). Mice were provided with water and food (Scientific Animal Food & Engineering, R150) ad libitum and kept on a 12-hour light/dark cycle at 22°C +/- 2°C with relative humidity of 55% +/-10%.

Mouse tissue collection, staining procedures, image acquisition and analysis

For intestinal whole-mount immunostaining (Bernier-Latmani and Petrova, 2016) pup intestines were dissected and fixed overnight in fixation buffer (0.5% paraformaldehyde (PFA), 15% picric acid, 1X PBS). After washing in 1X PBS, intestinal pieces were incubated with primary and secondary antibodies overnight, with washing all day after staining in wash buffer (0.3% Triton-X100, 1X PBS). After washing, samples were post-fixed with 4% PFA, 1X PBS overnight. All steps were performed at 4°C. After samples were cleared with RIMS buffer (Yang et al., 2014) and mounted for imaging. Primary antibodies are listed in Table S5 and were resuspended according to manufacturers’ recommendations when supplied lyophilized. Alexa Fluor 488, 555, and 647 fluorochrome-conjugated secondary antibodies (Invitrogen) were used for signal detection. Confocal images were obtained using a Zeiss LSM 880 microscope and ZEN acquisition software (Zeiss), using 20X (0.5 NA) or 40X (1.3 NA) objectives, at room temperature in RIMS. Images were analyzed using Imaris (Bitplane), Fiji (Schindelin et al., 2012), Photoshop (Adobe) and Affinity Photo 2 software. Quantification of confocal micrographs was performed on 4-7 individual image stacks with 1 to 4 villi per image, on average. Statistical tests performed are indicated in figure legends. Data distribution was assumed to be normal but this was not formally tested.

Intestinal fibroblast sorting

For intestinal fibroblast sorting (Bernier-Latmani et al., 2022), mice were sacrificed and the intestine was dissected and flushed with ice-cold PBS. Peyer’s patches were removed and the intestine was cut into 1 cm pieces, which were put in a 10 mM EDTA DMEM solution agitating at 37 °C for 30 min. to remove epithelial cells. The remaining tissue was digested thrice with Liberase TL (192.5 µg/mL, Roche) in DMEM (Gibco) containing 2% FBS, CaCl2 (2 mM) and 50ug/ml DNase I with constant stirring at 37 °C for 20 min and washed with medium. For fibroblast isolation from pups for RNAseq the cell suspension was incubated with labeled antibodies listed in Table S5 and the gating strategy is shown in (Fig. S1D). For isolation of GFP+ intestinal fibroblasts from adult Pdgfrb-GFP mice DAPI-, GFP+ cells were collected and the gating strategy is shown in (Fig. S1E). Cells were sorted with a BD FACSAria II (SORP) v8.0.1 cell sorter with BD FACSDiva software (BD Biosciences). After sorting, fibroblasts were collected in RLT Buffer (Qiagen) and snap frozen.

Pdgfrb-GFP fibroblast culture

50,000 Pdgfrb-GFP fibroblasts were plated per well on fibronectin-coated 96-well plates in RPMI (ThermoFisher) supplemented with 10% FBS and penicillin/streptomycin. Cells were allowed to attach and expand for 3 days, with media changes every 24 hours to remove non-adherent cells.

Isolation of intestinal eosinophils

Intestinal eosinophils were collected from intestinal single-cell suspensions from C57BL/6 adult mouse intestines with the same protocol as for intestinal fibroblasts. Eosinophils were isolated using Anti-Siglec-F MicroBeads (Miltenyi Biotec, #130-118-513) according to the manufacturer’s instructions. Briefly, single-cell suspensions were centrifuged at 300×g for 10 min at 4°C and resuspended in MACS buffer (PBS pH 7.2 supplemented with 0.5% BSA and 2 mM EDTA). Cells were incubated with Anti-Siglec-F MicroBeads for 10 min at 2–8°C. After washing with MACS buffer and centrifugation (300×g, 10 min), cells were resuspended and applied to MS columns placed in a magnetic field (MiniMACS Separator). Unlabeled cells were allowed to pass through. Columns were washed three times with buffer, removed from the magnetic field, and SIGLECF+ cells were eluted. To increase purity, the positive fraction was subjected to a second magnetic separation step over a new column as recommended by the manufacturer. Purity of isolated eosinophils was confirmed by staining for CCR3 and SIGLECF (Fig. S1F, Table S5) and analyzing on a LSRFortessa Cell Analyzer (BD Biosciences). Isolated SIGLECF+ cells were counted and immediately used for co-culture experiments.

Transwell co-culture and TGFβ treatment of intestinal fibroblasts

After Pdgfrb-GFP fibroblast attachment (day 3), culture medium was replaced and 96-well transwell inserts were placed into each well. SIGLECF+ eosinophils were seeded into the upper chamber of the insert (50,000 cells per insert) to allow indirect co-culture via soluble factors. Other wells with seeded fibroblasts received BSA (20ng/ml) or recombinant TGFβ1 (PeproTech, 100-21-2UG, 10ng/ml) or TGFβ2 (ThermoFisher, 100-35B-10UG, 10ng/ml) with or without SB431542 (Tocris, 5 µM). After co-culture or TGFβ treatment Pdgfrb-GFP fibroblasts were maintained for 72 hours in RPMI supplemented with 10% FBS and penicillin/streptomycin before fixation in 4% PFA and antibody staining.

Bulk RNAseq

RNA was extracted from sorted fibroblasts using the RNAeasy Mini kit (Qiagen) according to manufacturer’s instructions. RNA-seq libraries were prepared from 100 ng of total RNA using the TruSeq stranded mRNA library construction kit (Illumina) using manufacturer’s recommendations. The samples were sequenced on a HiSeq 2500 device (Illumina) with 125 single-end reads. Image analysis and base calling were conducted by the HiSeq Control software.

Bioinformatic analysis

Raw sequence data (.bcl files) generated from Illumina HiSeq was converted into fastq files and de-multiplexed using Illumina’s bcl2fastq v.2.2 software. One mismatch was allowed for index sequence identification. We used the kallisto pseudo-aligner (v. 0.44.0, with default settings plus bias correction)(Bray et al., 2016) to map the reads to the mm10 mouse reference genome (Ensembl release GRCm38.p6). Raw counts per transcripts were imported into R (v. 4.2.2) and aggregated at the gene level using the tximport package (v. 1.26.1)(Soneson et al., 2015). Genes with at least 0.33 cpm in at least 3 samples were retained (n=15238). Because a principal component analysis revealed a potential batch effect, we estimated surrogate variables using the svaseq function of the SVA package (v. 3.46.0). Differential expression analysis between KO and WT samples was performed with DESeq2 (Love et al., 2014), with 2 surrogate variables included in the design model. Genes with Benjamini-Hochberg adjusted p-value < 0.05 were considered as significantly differentially expressed. Functional analysis was conducted using Gene Set Enrichment Analysis (Subramanian et al., 2005) with the GSEA function of the clusterProfiler package (v. 4.6.2) (Wu et al., 2021), using as input the sorted Wald statistic of the genes with at least 1000 average normalized counts (n=5256) and the Hallmark gene set collection (Liberzon et al., 2015) accessed using the msigdbr package (v. 7.5.1(Liberzon et al., 2015; Subramanian et al., 2005). Gene sets with Benjamini-Hochberg adjusted p-value < 0.05 were considered as significantly enriched.

Differential gene expression analysis in lung versus gut eosinophils (Wen et al, 2012, GEO accession GSE33807) and in CD22+ gut versus α4β7-CD22- eosinophils (Kutyavin et al, 2024, GEO accession GSE236132) were performed using the limma package (v. 3.50.3) (Ritchie et al., 2015).

Block face scanning electron microscopy

Ten-week-old adult mice were perfused, via the heart, with a buffered mix of 2.5 % glutaraldehyde and 2.0 % paraformaldehyde in 0.1 M phosphate buffer (pH 7.4). The small intestine was removed and embedded in 5% agarose, and 80 µm thick, transverse sections, cut with a vibratome. The sections were post-fixed in potassium ferrocyanide (1.5%) and osmium (2%), then stained with thiocarbohydrazide (1%) followed by osmium tetroxide (2%). They were then stained overnight in uranyl acetate (1%), washed in distilled water at 50 °C, before being stained with lead aspartate at the same temperature. They were finally dehydrated in increasing concentrations of ethanol and then embedded in Spurr’s resin and hardened at 65 °C for 24 h between glass slides. Resin trimming with a glass knife produced a small (approximately 300 × 300 µm) block that was then mounted inside a scanning electron microscope (Zeiss Merlin, Zeiss NTS) holding a block face cutting microtome (3View, Gatan). Layers of resin, 50 nm thick, were cut from the block surface, and sequential images collected after each layer was removed. An acceleration voltage of the 1.7 kV was used with a pixel size of 7 nm with a dwell time of 1 µs. Series nearly aligned images were collected and aligned in the FIJI imaging software (www.fiji.sc). Segmentation of different cells and structures and video generation were carried out with FIJI software running the TrakEM2 plugin. Cells were counted as in direct contact when eosinophil membrane was in direct contact with membranes of other cells.

Statistical analyses and data presentation

GraphPad Prism 10 and 11 were used for statistical analyses and presentation of quantitative data. Two-tailed, unpaired Student’s t test was used for comparing two samples. One-way ANOVA was used to compare more than two conditions. When a P value of ANOVA was lower than 0.05, post hoc analysis (Tukey’s multiple comparisons) was performed. All values shown as mean ± SD. P values were indicated as follows: *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001.

Online supplemental material

Supplementary materials include 3 figures with legends (Fig. S1-3), 5 tables (Table S1-5) and 1 video (Video 1). Fig. S1 contains image analysis and FACS plots with additional analysis related to Fig. 1, 2, 4 and 5. Fig. S2 contains functional data from pup intestines after TGFβ and VEGFR3 blockade related to Fig. 5. Fig. S3 contains a scheme summarizing the main findings of this study. Table S1 contains differential gene expression of re-analyzed bulk RNAseq data from lung and intestinal eosinophils (Wen et al., 2012). Table S2 contains differential gene expression of re-analyzed bulk RNAseq data from intestinal CD22+ and α4β7-CD22- eosinophils (Kutyavin et al., 2024). Table S3 contains differential gene expression analysis and Table S4 contains gene set enrichment analysis from bulk RNAseq data of intestinal fibroblasts from control Balb/c and eosinophil-deficient ΔdblGata1 pups. Table S5 contains antibodies used in this study. Video 1 contains a 3D reconstruction of intestinal villus cells from a block face scanning electron microscopy image related to Fig. 2E.

Supplementary Material

Video 1
Download video file (850.3KB, mp4)
Figure S1
Figure S2
Figure S3
Table S1
Table S2
Table S3
Table S4
Table S5
Supplementary Figure Legends

eTOC summary.

Eosinophils are associated with infection and pathological fibrosis. However, healthy small intestinal villi harbor dense populations of these granulocytes. Here, Petrova et al., show that eosinophils convert PDGFRβ+ fibroblasts to specialized villus smooth muscle cells, driving a developmental program for postnatal intestinal remodeling and maturation.

Acknowledgements

We thank Christer Betsholtz and Michael Vanlandewijck (Karolinska Institute, Stockholm, Sweden) for providing samples from Pdgfrb-GFP mice, Amrita Manchala, Julia Baldwin and Sina Nassiri for participation in initial stages of this work, and Céline Beauverd for mouse genotyping and colony maintenance. The UNIL Animal, Cellular Imaging, Genomic Technologies, Mouse Pathology and Flow Cytometry Facilities and EPFL Bioelectron Microscopy Core Facility are gratefully acknowledged. Illustrations were generated with Biorender.

This work was supported by grants from the Swiss National Science Foundation (310030_197878) to TVP and the Novartis Foundation for Medical-Biological Research and Olga Mayenfisch Foundation to JBL.

Footnotes

Author contributions

JBL, KDK, TB, YM and LS performed experiments and analyzed data. TW performed bioinformatics analysis. KS and NLH provided reagents. JBL prepared the figures. JBL and TVP conceived the study, designed and supervised all experiments and wrote the manuscript. All authors read and reviewed the manuscript.

Declaration of interests

The authors declare no conflict of interest.

Data availability

The data underlying our re-analysis of previously published bulk RNAseq data in Fig. 4B (Wen et al., 2012) and Fig. 4C (Kutyavin et al., 2024) are publicly available (GSE33807 and GSE236132) and also provided as Table S1 and Table S2, respectively. The data underlying bulk RNAseq data in Fig. D-F are publicly available (GSE289872) and provided as Table S3 and S4. All other data are available from the corresponding authors upon reasonable request.

References

  1. Aceves SS, Newbury RO, Dohil R, Bastian JF, Broide DH. Esophageal remodeling in pediatric eosinophilic esophagitis. J Allergy Clin Immunol. 2007;119:206–212. doi: 10.1016/j.jaci.2006.10.016. [DOI] [PubMed] [Google Scholar]
  2. Arnold IC, Artola-Borán M, Tallón de Lara P, Kyburz A, Taube C, Ottemann K, van den Broek M, Yousefi S, Simon HU, Müller A. Eosinophils suppress Th1 responses and restrict bacterially induced gastrointestinal inflammation. J Exp Med. 2018;215:2055–2072. doi: 10.1084/jem.20172049. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Bernier-Latmani J, Cisarovsky C, Demir CS, Bruand M, Jaquet M, Davanture S, Ragusa S, Siegert S, Dormond O, Benedito R, Radtke F, et al. DLL4 promotes continuous adult intestinal lacteal regeneration and dietary fat transport. J Clin Invest. 2015;125:4572–4586. doi: 10.1172/JCI82045. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Bernier-Latmani J, González-Loyola A, Petrova TV. Mechanisms and functions of intestinal vascular specialization. J Exp Med. 2024;221 doi: 10.1084/jem.20222008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bernier-Latmani J, Mauri C, Marcone R, Renevey F, Durot S, He L, Vanlandewijck M, Maclachlan C, Davanture S, Zamboni N, Knott GW, et al. ADAMTS18+ villus tip telocytes maintain a polarized VEGFA signaling domain and fenestrations in nutrient-absorbing intestinal blood vessels. Nat Commun. 2022;13:3983. doi: 10.1038/s41467-022-31571-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Bernier-Latmani J, Petrova TV. High-resolution 3D analysis of mouse small-intestinal stroma. Nat Protoc. 2016;11:1617–1629. doi: 10.1038/nprot.2016.092. [DOI] [PubMed] [Google Scholar]
  7. Bernier-Latmani J, Petrova TV. Intestinal lymphatic vasculature: structure, mechanisms and functions. Nat Rev Gastroenterol Hepatol. 2017;14:510–526. doi: 10.1038/nrgastro.2017.79. [DOI] [PubMed] [Google Scholar]
  8. Bray NL, Pimentel H, Melsted P, Pachter L. Near-optimal probabilistic RNA-seq quantification. Nat Biotechnol. 2016;34:525–527. doi: 10.1038/nbt.3519. [DOI] [PubMed] [Google Scholar]
  9. Choe K, Jang JY, Park I, Kim Y, Ahn S, Park D-Y, Hong Y-K, Alitalo K, Koh GY, Kim P. Intravital imaging of intestinal lacteals unveils lipid drainage through contractility. J Clin Invest. 2015;125:4042–4052. doi: 10.1172/JCI76509. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Cuervo H, Pereira B, Nadeem T, Lin M, Lee F, Kitajewski J, Lin C-S. PDGFRβ-P2A-CreERT2 mice: a genetic tool to target pericytes in angiogenesis. Angiogenesis. 2017;20:655–662. doi: 10.1007/s10456-017-9570-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Diny NL, Schonfeldova B, Shapiro M, Winder ML, Varsani-Brown S, Stockinger B. The aryl hydrocarbon receptor contributes to tissue adaptation of intestinal eosinophils in mice. J Exp Med. 2022;219 doi: 10.1084/jem.20210970. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Ewing-Crystal NA, Mroz NM, Larpthaveesarp A, Lizama CO, Pennington R, Chiaranunt P, Dennis JI, Chang AA, Merrill ED, Caryotakis SE, Kirthivasan N, et al. Dynamic fibroblast–immune interactions shape recovery after brain injury. Nature. 2025;646:934–944. doi: 10.1038/s41586-025-09449-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Fallegger A, Priola M, Artola-Borán M, Núñez NG, Wild S, Gurtner A, Becher B, Yousefi S, Simon H-U, Arnold IC, Müller A. TGF-β production by eosinophils drives the expansion of peripherally induced neuropilin− RORγt+ regulatory T-cells during bacterial and allergen challenge. Mucosal Immunol. 2022;15:504–514. doi: 10.1038/s41385-022-00484-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Fettrelet T, Gigon L, Karaulov A, Yousefi S, Simon HU. The Enigma of Eosinophil Degranulation. Int J Mol Sci. 2021;22 doi: 10.3390/ijms22137091. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Frangogiannis N. Transforming growth factor-β in tissue fibrosis. J Exp Med. 2020;217:e20190103. doi: 10.1084/jem.20190103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Furuta GT, Katzka DA. Eosinophilic Esophagitis. N Engl J Med. 2015;373:1640–1648. doi: 10.1056/NEJMra1502863. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Gehart H, Clevers H. Tales from the crypt: new insights into intestinal stem cells. Nature Reviews Gastroenterology & Hepatology. 2019;16:19–34. doi: 10.1038/s41575-018-0081-y. [DOI] [PubMed] [Google Scholar]
  18. Gigon L, Fettrelet T, Yousefi S, Simon D, Simon H-U. Eosinophils from A to Z. Allergy. 2023;78:1810–1846. doi: 10.1111/all.15751. [DOI] [PubMed] [Google Scholar]
  19. Gong S, Zheng C, Doughty ML, Losos K, Didkovsky N, Schambra UB, Nowak NJ, Joyner A, Leblanc G, Hatten ME, Heintz N. A gene expression atlas of the central nervous system based on bacterial artificial chromosomes. Nature. 2003;425:917–925. doi: 10.1038/nature02033. [DOI] [PubMed] [Google Scholar]
  20. Gouon-Evans V, Rothenberg ME, Pollard JW. Postnatal mammary gland development requires macrophages and eosinophils. Development. 2000;127:2269–2282. doi: 10.1242/dev.127.11.2269. [DOI] [PubMed] [Google Scholar]
  21. Gurtner A, Borrelli C, Gonzalez-Perez I, Bach K, Acar IE, Núñez NG, Crepaz D, Handler K, Vu VP, Lafzi A, Stirm K, et al. Active eosinophils regulate host defence and immune responses in colitis. Nature. 2023a;615:151–157. doi: 10.1038/s41586-022-05628-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Gurtner A, Crepaz D, Arnold IC. Emerging functions of tissue-resident eosinophils. J Exp Med. 2023b;220:e20221435. doi: 10.1084/jem.20221435. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Hargreaves M, Spriet LL. Skeletal muscle energy metabolism during exercise. Nature Metabolism. 2020;2:817–828. doi: 10.1038/s42255-020-0251-4. [DOI] [PubMed] [Google Scholar]
  24. Hong SP, Yang MJ, Cho H, Park I, Bae H, Choe K, Suh SH, Adams RH, Alitalo K, Lim D, Koh GY. Distinct fibroblast subsets regulate lacteal integrity through YAP/TAZ-induced VEGF-C in intestinal villi. Nat Commun. 2020;11:4102. doi: 10.1038/s41467-020-17886-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Hu S, Mahadevan A, Elysee IF, Choi J, Souchet NR, Bae GH, Taboada AK, Sanketi B, Duhamel GE, Sevier CS, Tao G, et al. The asymmetric Pitx2 gene regulates gut muscular-lacteal development and protects against fatty liver disease. Cell Rep. 2021;37:110030. doi: 10.1016/j.celrep.2021.110030. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Huycke TR, Miller BM, Gill HK, Nerurkar NL, Sprinzak D, Mahadevan L, Tabin CJ. Genetic and Mechanical Regulation of Intestinal Smooth Muscle Development. Cell. 2019;179:90–105.:e121. doi: 10.1016/j.cell.2019.08.041. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Ignacio A, Shah K, Bernier-Latmani J, Köller Y, Coakley G, Moyat M, Hamelin R, Armand F, Wong NC, Ramay H, Thomson CA, et al. Small intestinal resident eosinophils maintain gut homeostasis following microbial colonization. Immunity. 2022;55:1250–1267.:e1212. doi: 10.1016/j.immuni.2022.05.014. [DOI] [PubMed] [Google Scholar]
  28. Jung Y, Wen T, Mingler MK, Caldwell JM, Wang YH, Chaplin DD, Lee EH, Jang MH, Woo SY, Seoh JY, Miyasaka M, et al. IL-1β in eosinophil-mediated small intestinal homeostasis and IgA production. Mucosal Immunol. 2015;8:930–942. doi: 10.1038/mi.2014.123. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Kariyawasam HH, Robinson DS. The role of eosinophils in airway tissue remodelling in asthma. Curr Opin Immunol. 2007;19:681–686. doi: 10.1016/j.coi.2007.07.021. [DOI] [PubMed] [Google Scholar]
  30. Kim KE, Sung H-K, Koh GY. Lymphatic development in mouse small intestine. Dev Dyn. 2007;236:2020–2025. doi: 10.1002/dvdy.21200. [DOI] [PubMed] [Google Scholar]
  31. Kutyavin V, Korn LL, Medzhitov R. Nutrient-derived signals regulate eosinophil adaptation to the small intestine. Proceedings of the National Academy of Sciences. 2024;121:e2316446121. doi: 10.1073/pnas.2316446121. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Le VQ, Zhao B, Ramesh S, Toohey C, DeCosta A, Mintseris J, Liu X, Gygi S, Springer TA. A specialized integrin-binding motif enables proTGF-β2 activation by integrin αVβ6 but not αVβ8. Proc Natl Acad Sci U S A. 2023;120:e2304874120. doi: 10.1073/pnas.2304874120. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Lee JS. Contraction of villi and fluid transport in dog jejunal mucosa in-vitro. Am J Physiol. 1971;221:488–495. doi: 10.1152/ajplegacy.1971.221.2.488. [DOI] [PubMed] [Google Scholar]
  34. Levéen P, Larsson J, Ehinger M, Cilio CM, Sundler M, Sjöstrand LJ, Holmdahl R, Karlsson S. Induced disruption of the transforming growth factor beta type II receptor gene in mice causes a lethal inflammatory disorder that is transplantable. Blood. 2002;100:560–568. doi: 10.1182/blood.v100.2.560. [DOI] [PubMed] [Google Scholar]
  35. Li Y, Liu S, Zhou K, Wang Y, Chen Y, Hu W, Li S, Li H, Wang Y, Wang Q, He D, et al. Neuromedin U programs eosinophils to promote mucosal immunity of the small intestine. Science. 2023;381:1189–1196. doi: 10.1126/science.ade4177. [DOI] [PubMed] [Google Scholar]
  36. Liberzon A, Birger C, Thorvaldsdóttir H, Ghandi M, Mesirov JP, Tamayo P. The Molecular Signatures Database (MSigDB) hallmark gene set collection. Cell Syst. 2015;1:417–425. doi: 10.1016/j.cels.2015.12.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Loffredo LF, Coden ME, Jeong BM, Walker MT, Anekalla KR, Doan TC, Rodriguez R, Browning M, Nam K, Lee JJ, Abdala-Valencia H, et al. Eosinophil accumulation in postnatal lung is specific to the primary septation phase of development. Scientific Reports. 2020;10:4425. doi: 10.1038/s41598-020-61420-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Love MI, Huber W, Anders S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 2014;15:550. doi: 10.1186/s13059-014-0550-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Madison B, Braunstein K, Kuizon E, Portman K, Qiao X, Gumucio D. Epithelial hedgehog signals pattern the intestinal crypt-villus axis. Development. 2005;132:279–289. doi: 10.1242/dev.01576. [DOI] [PubMed] [Google Scholar]
  40. Massagué J, Sheppard D. TGF-β signaling in health and disease. Cell. 2023;186:4007–4037. doi: 10.1016/j.cell.2023.07.036. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. McBryan J, Howlin J, Kenny PA, Shioda T, Martin F. ERalpha-CITED1 co-regulated genes expressed during pubertal mammary gland development: implications for breast cancer prognosis. Oncogene. 2007;26:6406–6419. doi: 10.1038/sj.onc.1210468. [DOI] [PubMed] [Google Scholar]
  42. Mishra A, Hogan SP, Lee JJ, Foster PS, Rothenberg ME. Fundamental signals that regulate eosinophil homing to the gastrointestinal tract. J Clin Invest. 1999;103:1719–1727. doi: 10.1172/JCI6560. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Muhl L, Genové G, Leptidis S, Liu J, He L, Mocci G, Sun Y, Gustafsson S, Buyandelger B, Chivukula IV, Segerstolpe Å, et al. Single-cell analysis uncovers fibroblast heterogeneity and criteria for fibroblast and mural cell identification and discrimination. Nat Commun. 2020;11:3953. doi: 10.1038/s41467-020-17740-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Muzumdar MD, Tasic B, Miyamichi K, Li L, Luo L. A global double-fluorescent Cre reporter mouse. Genesis. 2007;45:593–605. doi: 10.1002/dvg.20335. [DOI] [PubMed] [Google Scholar]
  45. Nurmi H, Saharinen P, Zarkada G, Zheng W, Robciuc MR, Alitalo K. VEGF-C is required for intestinal lymphatic vessel maintenance and lipid absorption. EMBO Mol Med. 2015;7:1418–1425. doi: 10.15252/emmm.201505731. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Papi A, Brightling C, Pedersen SE, Reddel HK. Asthma. The Lancet. 2018;391:783–800. doi: 10.1016/S0140-6736(17)33311-1. [DOI] [PubMed] [Google Scholar]
  47. Plasari G, Calabrese A, Dusserre Y, Gronostajski RM, McNair A, Michalik L, Mermod N. Nuclear factor I-C links platelet-derived growth factor and transforming growth factor beta1 signaling to skin wound healing progression. Mol Cell Biol. 2009;29:6006–6017. doi: 10.1128/MCB.01921-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Powell DW, Mifflin RC, Valentich JD, Crowe SE, Saada JI, West AB. Myofibroblasts. II. Intestinal subepithelial myofibroblasts. American Journal of Physiology-Cell Physiology. 1999;277:C183–C201. doi: 10.1152/ajpcell.1999.277.2.C183. [DOI] [PubMed] [Google Scholar]
  49. Pytowski B, Goldman J, Persaud K, Wu Y, Witte L, Hicklin DJ, Skobe M, Boardman KC, Swartz MA. Complete and specific inhibition of adult lymphatic regeneration by a novel VEGFR-3 neutralizing antibody. J Natl Cancer Inst. 2005;97:14–21. doi: 10.1093/jnci/dji003. [DOI] [PubMed] [Google Scholar]
  50. Ritchie ME, Phipson B, Wu D, Hu Y, Law CW, Shi W, Smyth GK. limma powers differential expression analyses for RNA-sequencing and microarray studies. Nucleic Acids Res. 2015;43:e47. doi: 10.1093/nar/gkv007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Roci I, Kim J, de Korodi K, Wyss T, Bernier-Latmani J, Arroz-Madeira S, Bovay E, Schoofs H, Mäkinen T, Noël A, Petrova TV. Aquaporin-1 sustains lymphangiogenic responses in inflammatory microenvironments. bioRxiv. 2025:2025.2003.2005.641652. doi: 10.1084/jem.20250506. [DOI] [PubMed] [Google Scholar]
  52. Sanketi BD, Mantri M, Huang L, Tavallaei MA, Hu S, Wang MFZ, De Vlaminck I, Kurpios NA. Villus myofibroblasts are developmental and adult progenitors of mammalian gut lymphatic musculature. Dev Cell. 2024;59:1159–1174.:e1155. doi: 10.1016/j.devcel.2024.03.005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez J-Y, et al. Fiji: an open-source platform for biological-image analysis. Nat Methods. 2012;9:676–682. doi: 10.1038/nmeth.2019. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Shyer AE, Tallinen T, Nerurkar NL, Wei Z, Gil ES, Kaplan DL, Tabin CJ, Mahadevan L. Villification: how the gut gets its villi. Science. 2013;342:212–218. doi: 10.1126/science.1238842. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Soneson C, Love MI, Robinson MD. Differential analyses for RNA-seq: transcript-level estimates improve gene-level inferences. F1000Res. 2015;4:1521. doi: 10.12688/f1000research.7563.2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Spencer LA, Bonjour K, Melo RCN, Weller PF. Eosinophil Secretion of Granule-Derived Cytokines. Frontiers in Immunology. 2014;5 doi: 10.3389/fimmu.2014.00496. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Subramanian A, Tamayo P, Mootha VK, Mukherjee S, Ebert BL, Gillette MA, Paulovich A, Pomeroy SL, Golub TR, Lander ES, Mesirov JP. Gene set enrichment analysis: A knowledge-based approach for interpreting genome-wide expression profiles. Proc Natl Acad Sci U S A. 2005;102:15545–15550. doi: 10.1073/pnas.0506580102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Suh SH, Choe K, Hong SP, Jeong S-h, Mäkinen T, Kim KS, Alitalo K, Surh CD, Koh GY, Song J-H. Gut microbiota regulates lacteal integrity by inducing VEGF-C in intestinal villus macrophages. EMBO Rep. 2019;20:e46927. doi: 10.15252/embr.201846927. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Takemura N, Kurashima Y, Mori Y, Okada K, Ogino T, Osawa H, Matsuno H, Aayam L, Kaneto S, Park EJ, Sato S, et al. Eosinophil depletion suppresses radiation-induced small intestinal fibrosis. Sci Transl Med. 2018;10:eaan0333. doi: 10.1126/scitranslmed.aan0333. [DOI] [PubMed] [Google Scholar]
  60. Tammela T, Zarkada G, Wallgard E, Murtomaki A, Suchting S, Wirzenius M, Waltari M, Hellstrom M, Schomber T, Peltonen R, Freitas C, et al. Blocking VEGFR-3 suppresses angiogenic sprouting and vascular network formation. Nature. 2008;454:656–660. doi: 10.1038/nature07083. [DOI] [PubMed] [Google Scholar]
  61. Tso P, Bernier-Latmani J, Petrova TV, Liu M. Transport functions of intestinal lymphatic vessels. Nature Reviews Gastroenterology & Hepatology. 2024 doi: 10.1038/s41575-024-00996-z. [DOI] [PubMed] [Google Scholar]
  62. Walton KD, Whidden M, Kolterud Å, Shoffner SK, Czerwinski MJ, Kushwaha J, Parmar N, Chandhrasekhar D, Freddo AM, Schnell S, Gumucio DL. Villification in the mouse: Bmp signals control intestinal villus patterning. Development. 2016;143:427–436. doi: 10.1242/dev.130112. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Wang G, Jacquet L, Karamariti E, Xu Q. Origin and differentiation of vascular smooth muscle cells. J Physiol. 2015;593:3013–3030. doi: 10.1113/JP270033. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Wang W-L, Kasamatsu J, Joshita S, Gilfillan S, Di Luccia B, Panda SK, Kim D-H, Desai P, Bando JK, Huang SC-C, Yomogida K, et al. The aryl hydrocarbon receptor instructs the immunomodulatory profile of a subset of Clec4a4+ eosinophils unique to the small intestine. Proceedings of the National Academy of Sciences. 2022;119:e2204557119. doi: 10.1073/pnas.2204557119. [DOI] [PMC free article] [PubMed] [Google Scholar]
  65. Wen T, Mingler MK, Blanchard C, Wahl B, Pabst O, Rothenberg ME. The Pan-B Cell Marker CD22 Is Expressed on Gastrointestinal Eosinophils and Negatively Regulates Tissue Eosinophilia. J Immunol. 2012;188:1075–1082. doi: 10.4049/jimmunol.1102222. [DOI] [PMC free article] [PubMed] [Google Scholar]
  66. Womack WA, Barrowman JA, Graham WH, Benoit JN, Kvietys PR, Granger DN. Quantitative assessment of villous motility. Am J Physiol. 1987;252:G250–G256. doi: 10.1152/ajpgi.1987.252.2.G250. [DOI] [PubMed] [Google Scholar]
  67. Womack WA, Tygart PK, Mailman D, Kvietys PR, Granger DN. Villous motility - relationship to lymph-flow and blood-flow in the dog jejunum. Gastroenterology. 1988;94:977–983. doi: 10.1016/0016-5085(88)90556-2. [DOI] [PubMed] [Google Scholar]
  68. Wu D, Molofsky AB, Liang HE, Ricardo-Gonzalez RR, Jouihan HA, Bando JK, Chawla A, Locksley RM. Eosinophils sustain adipose alternatively activated macrophages associated with glucose homeostasis. Science. 2011;332:243–247. doi: 10.1126/science.1201475. [DOI] [PMC free article] [PubMed] [Google Scholar]
  69. Wu T, Hu E, Xu S, Chen M, Guo P, Dai Z, Feng T, Zhou L, Tang W, Zhan L, Fu X, et al. clusterProfiler 4.0: A universal enrichment tool for interpreting omics data. Innovation (Camb) 2021;2:100141. doi: 10.1016/j.xinn.2021.100141. [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Yang B, Treweek JB, Kulkarni RP, Deverman BE, Chen C-K, Lubeck E, Shah S, Cai L, Gradinaru V. Single-cell phenotyping within transparent intact tissue through whole-body clearing. Cell. 2014;158:945–958. doi: 10.1016/j.cell.2014.07.017. [DOI] [PMC free article] [PubMed] [Google Scholar]
  71. Yu C, Cantor AB, Yang H, Browne C, Wells RA, Fujiwara Y, Orkin SH. Targeted deletion of a high-affinity GATA-binding site in the Gata-1 promoter leads to selective loss of the eosinophil lineage in vivo. J Exp Med. 2002;195:1387–1395. doi: 10.1084/jem.20020656. [DOI] [PMC free article] [PubMed] [Google Scholar]
  72. Zhang F, Zarkada G, Han J, Li J, Dubrac A, Ola R, Genet G, Boyé K, Michon P, Künzel SE, Camporez JP, et al. Lacteal junction zippering protects against diet-induced obesity. Science. 2018;361:599–603. doi: 10.1126/science.aap9331. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Video 1
Download video file (850.3KB, mp4)
Figure S1
EMS215599-supplement-Figure_S1.tiff (3.1MB, tiff)
Figure S2
EMS215599-supplement-Figure_S2.tiff (3.8MB, tiff)
Figure S3
EMS215599-supplement-Figure_S3.tiff (887.6KB, tiff)
Table S1
EMS215599-supplement-Table_S1.xlsx (8.9MB, xlsx)
Table S2
EMS215599-supplement-Table_S2.xlsx (2.5MB, xlsx)
Table S3
EMS215599-supplement-Table_S3.xlsx (510KB, xlsx)
Table S4
EMS215599-supplement-Table_S4.xlsx (35.2KB, xlsx)
Table S5
EMS215599-supplement-Table_S5.xlsx (11.2KB, xlsx)
Supplementary Figure Legends
EMS215599-supplement-Supplementary_Figure_Legends.pdf (152.2KB, pdf)

Data Availability Statement

The data underlying our re-analysis of previously published bulk RNAseq data in Fig. 4B (Wen et al., 2012) and Fig. 4C (Kutyavin et al., 2024) are publicly available (GSE33807 and GSE236132) and also provided as Table S1 and Table S2, respectively. The data underlying bulk RNAseq data in Fig. D-F are publicly available (GSE289872) and provided as Table S3 and S4. All other data are available from the corresponding authors upon reasonable request.

RESOURCES