Lymphatic vessels interact dynamically with the hair follicle stem cell niche during skin regeneration in vivo
2019; Springer Nature; Volume: 38; Issue: 19 Linguagem: Inglês
10.15252/embj.2019101688
ISSN1460-2075
AutoresDaniel Peña-Jiménez, Sílvia Fontenete, Diego Megı́as, Coral Fustero‐Torre, Osvaldo Graña‐Castro, Donatello Castellana, Robert Loewe, Mirna Pérez‐Moreno,
Tópico(s)Wound Healing and Treatments
ResumoArticle2 September 2019Open Access Transparent process Lymphatic vessels interact dynamically with the hair follicle stem cell niche during skin regeneration in vivo Daniel Peña-Jimenez Daniel Peña-Jimenez Epithelial Cell Biology Group, Cancer Cell Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Search for more papers by this author Silvia Fontenete Silvia Fontenete Epithelial Cell Biology Group, Cancer Cell Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Section of Cell Biology and Physiology, Department of Biology, University of Copenhagen, Copenhagen, Denmark Search for more papers by this author Diego Megias Diego Megias Confocal Microscopy Core Unit, Biotechnology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Search for more papers by this author Coral Fustero-Torre Coral Fustero-Torre Bioinformatics Unit, Structural Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Search for more papers by this author Osvaldo Graña-Castro Osvaldo Graña-Castro Bioinformatics Unit, Structural Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Search for more papers by this author Donatello Castellana Donatello Castellana Epithelial Cell Biology Group, Cancer Cell Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Center for Cooperative Research Biosciences (CIC bioGUNE), Derio Bizkaia, Spain Search for more papers by this author Robert Loewe Robert Loewe Department of Dermatology, Medical University of Vienna, Vienna, Austria Search for more papers by this author Mirna Perez-Moreno Corresponding Author Mirna Perez-Moreno [email protected] orcid.org/0000-0002-0170-6406 Epithelial Cell Biology Group, Cancer Cell Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Section of Cell Biology and Physiology, Department of Biology, University of Copenhagen, Copenhagen, Denmark Search for more papers by this author Daniel Peña-Jimenez Daniel Peña-Jimenez Epithelial Cell Biology Group, Cancer Cell Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Search for more papers by this author Silvia Fontenete Silvia Fontenete Epithelial Cell Biology Group, Cancer Cell Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Section of Cell Biology and Physiology, Department of Biology, University of Copenhagen, Copenhagen, Denmark Search for more papers by this author Diego Megias Diego Megias Confocal Microscopy Core Unit, Biotechnology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Search for more papers by this author Coral Fustero-Torre Coral Fustero-Torre Bioinformatics Unit, Structural Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Search for more papers by this author Osvaldo Graña-Castro Osvaldo Graña-Castro Bioinformatics Unit, Structural Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Search for more papers by this author Donatello Castellana Donatello Castellana Epithelial Cell Biology Group, Cancer Cell Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Center for Cooperative Research Biosciences (CIC bioGUNE), Derio Bizkaia, Spain Search for more papers by this author Robert Loewe Robert Loewe Department of Dermatology, Medical University of Vienna, Vienna, Austria Search for more papers by this author Mirna Perez-Moreno Corresponding Author Mirna Perez-Moreno [email protected] orcid.org/0000-0002-0170-6406 Epithelial Cell Biology Group, Cancer Cell Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain Section of Cell Biology and Physiology, Department of Biology, University of Copenhagen, Copenhagen, Denmark Search for more papers by this author Author Information Daniel Peña-Jimenez1,‡, Silvia Fontenete1,2,‡, Diego Megias3, Coral Fustero-Torre4, Osvaldo Graña-Castro4, Donatello Castellana1,5, Robert Loewe6 and Mirna Perez-Moreno *,1,2 1Epithelial Cell Biology Group, Cancer Cell Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain 2Section of Cell Biology and Physiology, Department of Biology, University of Copenhagen, Copenhagen, Denmark 3Confocal Microscopy Core Unit, Biotechnology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain 4Bioinformatics Unit, Structural Biology Programme, Spanish Cancer Research Centre (CNIO), Madrid, Spain 5Center for Cooperative Research Biosciences (CIC bioGUNE), Derio Bizkaia, Spain 6Department of Dermatology, Medical University of Vienna, Vienna, Austria ‡These authors contributed equally to this work *Corresponding author. Tel: +45 35 33 35 40; E-mail: [email protected] The EMBO Journal (2019)38:e101688https://doi.org/10.15252/embj.2019101688 [Correction added on 1 October 2019, after first online publication: the author affiliations have been updated.] See also: CY Kam & V Greco (October 2019) PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Lymphatic vessels are essential for skin fluid homeostasis and immune cell trafficking. Whether the lymphatic vasculature is associated with hair follicle regeneration is, however, unknown. Here, using steady and live imaging approaches in mouse skin, we show that lymphatic vessels distribute to the anterior permanent region of individual hair follicles, starting from development through all cycle stages and interconnecting neighboring follicles at the bulge level, in a stem cell-dependent manner. Lymphatic vessels further connect hair follicles in triads and dynamically flow across the skin. At the onset of the physiological stem cell activation, or upon pharmacological or genetic induction of hair follicle growth, lymphatic vessels transiently expand their caliber suggesting an increased tissue drainage capacity. Interestingly, the physiological caliber increase is associated with a distinct gene expression correlated with lymphatic vessel reorganization. Using mouse genetics, we show that lymphatic vessel depletion blocks hair follicle growth. Our findings point toward the lymphatic vasculature being important for hair follicle development, cycling, and organization, and define lymphatic vessels as stem cell niche components, coordinating connections at tissue-level, thus provide insight into their functional contribution to skin regeneration. Synopsis Like blood vasculature, lymphangiogenesis may have an impact on epithelial tissue development. Combined morphological profiling and genetic analyses now show structural and functional association between lymph vessels (LV) and the hair follicles (HF), providing new insights into principles of niche organization during skin regeneration and maintenance. LVs in the mouse back skin associate with HFs in a polarized manner, interconnecting HF triads throughout the hair cycle. Depletion of Wnt ligands in HF stem cells disrupts LV-HF association. LVs transiently increase their caliber at the onset of stem cell activation showing a distinct molecular signature. Genetic ablation of LVs leads to premature exit from HF growth. Introduction Lymphatic vessels (LV) play fundamental homeostatic functions including the balanced transport of fluids and macromolecules, the local coordination of immune responses, and immune cell trafficking to regional lymph nodes (Skobe & Detmar, 2000). After years of scientific discovery, much has been learned about the distinctive characteristics of LV, including the molecular markers prospero-related homeobox 1 (Prox1), lymphatic vessel endothelial hyaluronan receptor 1 (LYVE1), and podoplanin, as well as critical regulatory signals that govern their development and fundamental functions in tissues (Wang & Oliver, 2010; Yang & Oliver, 2014; Zheng et al, 2014; Potente & Makinen, 2017). In skin, LV are organized in structured polygonal patterns, consisting of one subcutaneous plexus, and a more superficial plexus located in the dermis, near the blood vessels (Braverman, 1989). Several studies have contributed to our understanding of the orderly organization of LV in the skin (Skobe & Detmar, 2000; Tripp et al, 2008). These studies have provided insight into the existence of branches of lymphatic capillaries that extend to the HF and drain into the subcutaneous collecting LV, presumably through connections of blind capillaries with the dermal papillae (dp) (Forbes, 1938), a condensate of dermal fibroblasts that provides a specialized microenvironment (Millar, 2002; Yang & Cotsarelis, 2010; Sennett & Rendl, 2012). LV may also facilitate the entry of immune cells to the HF epithelium, a source of chemokines that regulate the trafficking of epidermal Langerhans cells and dermal dendritic cells (Nagao et al, 2012), the distribution and differentiation of Langerhans cells (Wang et al, 2012), and the tropism of skin resident memory T cells (Adachi et al, 2015). However, despite the role of LV in facilitating immune cell trafficking to HF, less is known about their coordinated connections during HF cycling and functional implications. In adult skin, HF exhibits a lifetime polarized pattern of growth and regeneration across the tissue modulated by stimulatory and inhibitory signals (Plikus et al, 2011; Widelitz & Chuong, 2016). The cyclic regeneration of HF involves phases of growth (Anagen) via regression (Catagen) to relative quiescence (Telogen; Geyfman et al, 2015). The entry of resting HF into Anagen requires the activation of HFSC located in the HF bulge (Cotsarelis et al, 1990; Tumbar et al, 2004), and the expansion of their progenitors found in the secondary hair germ, giving rise to a new Anagen HF (Tumbar et al, 2004; Greco et al, 2009; Rompolas et al, 2012). Anagen HF grows until other instructive signals promote their regression giving rise to a new HF cycle. In past decades, a wealth of knowledge has yielded valuable insight into the role of major stimulatory and inhibitory signals in governing the orchestrated activation of the HF cycle (Blanpain & Fuchs, 2009; Lee & Tumbar, 2012; Plikus & Chuong, 2014), including local self-activation signals (Hsu et al, 2011), the contribution of other cells in the tissue macroenvironment (Brownell et al, 2011; Festa et al, 2011; Castellana et al, 2014; Rivera-Gonzalez et al, 2016; Ali et al, 2017), and long-range signaling waves across the skin (Plikus et al, 2011). Blood vessels have also been found associated around HF exhibiting a coordinated dynamic reorganization during HF cycling (Mecklenburg et al, 2000; Yano et al, 2001). Also, HFSC closely associates with a venule annulus (Xiao et al, 2013). The occurrence of angiogenesis, the growth of new capillaries from pre-existing blood vessels, has been observed during Anagen (Mecklenburg et al, 2000). The epidermal expression of the vascular endothelial growth factor A (VEGF-A; Detmar, 1996) induces perifollicular angiogenesis and sustains HF growth; conversely, inhibition of VEGF-A leads to a delay in HF growth accompanied by reductions in HF size and perifollicular vascularization (Yano et al, 2001). Overall, these results exposed that HF and blood vessels form a functional operative system. In contrast, less is known about the role of LV in regulating this process. Here, we show that lymphatic capillaries are novel components of the HFSC niche, coordinating HF connections at tissue-level and provide insight into their functional association to the HF cycle. Results Lymphatic capillaries distribute in the vicinity of HFSC in a polarized manner To investigate the association between LV and HFSC, we first defined the lymphatic distribution at HF in mouse back skin. To this end, we performed immunofluorescence analyses using antibodies to the lymphatic marker LYVE1 as well as to alpha smooth muscle actin (αSMA), enriched in the arrector pili muscle (apm). Lymphatic capillaries distributed in a polarized manner, aligned to the anterior side of the HF, opposite the distribution described for the apm (Fujiwara et al, 2011), ascending as blind capillaries along the HF permanent area toward the epidermis until the infundibulum (Fig 1A). This distribution was different from the one observed in the ear skin, which presented parallel lymphatic capillaries that were not associated with HF (Fig EV1A). In the back skin, lymphatic capillaries densely distributed to the anterior side of HF at Tenascin-C areas (Fig 1B), a glycoprotein of the extracellular matrix (ECM) enriched in the HF bulge (Tumbar et al, 2004). To interrogate the existence of a lymphatic association with both embryo and adult HFSC, we next determined the lymphatic distance to Lhx2+ SC (Fig 1C and D; Rhee et al, 2006) and CD34+ HFSC (Fig 1E and F; Blanpain et al, 2004), respectively. Immunofluorescence analyses of back skin sections of P5 and P12 mice (Fig 1C and D) and P49 mice (Fig 1E and F) revealed that lymphatic capillaries distributed to the proximity of HFSC, within a distance inferior of 3 μm, within the ratio expected for components of the HFSC niche (Beck et al, 2011). Figure 1. LV interact with HF in a functional HFSC niche-dependent manner Adult mouse back skin sections immunostained for LYVE1 (red), αSMA (green), and counterstained with DAPI (blue). n = 3–4 skin samples per mouse, n = 3–4 mice. Scale bar, 50 μm. LV, lymphatic vessels; apm, arrector pili muscle. Back skin sections of P55 mouse skin immunostained for LYVE1 (red), Tenascin-C (green), and counterstained with DAPI (blue). n = 3–4 skin samples per mouse, n = 3–4 mice. Scale bar, 50 μm. TNC, Tenascin-C. Adult mouse back skin sections from different postnatal (P) days immunostained for LYVE1 (red), Lhx2 (green), and counterstained with DAPI (blue). n = 3–4 skin samples per mouse, n = 3–4 mice. Scale bar, 50 μm. epi, epidermis; der, dermis. Histogram of the quantification of the LV distance to HFSC positive for Lhx2. n = 3–4 skin samples per mouse, n = 3–4 mice. Data represent the mean value ± SEM. ***P < 0.001 (Mann–Whitney U-test). Back skin sections of P49 mouse skin immunostained for LYVE1 (red), CD34 (green), and counterstained with DAPI (blue). n = 3–4 skin samples per mouse, n = 3–4 mice. Scale bar, 100 μm. epi, epidermis; der, dermis. White arrowheads denote the proximity of lymphatic capillaries to HFSC. Histogram of the quantification of the LV distance to HFSC positive for CD34. n = 3–4 skin samples per mouse, n = 3–4 mice. C, Catagen; Te, early Telogen; Tm, mid-Telogen; Tl, late Telogen; A, Anagen. Data represent the mean value ± SEM (Kruskal–Wallis test, Tukey's test). Back skin sections of K15CrePR+/T; WlsΔ/Δ and Control K15CrePR+/+; Wlsflox/flox mice treated with mifepristone from P7 during 12 weeks (P90), immunostained for LYVE1 (red), and counterstained with DAPI (blue). n = 3–4 skin samples per mouse, n = 3–4 mice. Scale bar, 100 μm. epi, epidermis; der, dermis. Histogram of the quantification of the LV angle relative to the epidermis. n = 3–4 skin samples per mouse, n = 3–4 mice. Data represent the mean value ± SEM. ***P < 0.001 (Mann–Whitney U-test). Download figure Download PowerPoint Click here to expand this figure. Figure EV1. Distribution of LV in the ear skin and validation of the reductions of Wls expression in the K15CrePR+/T; WlsΔ/Δ mouse model Adult ear skin sections immunostained for LYVE1 (red) and counterstained with DAPI (blue). Scale bar, 50 μm. LV, lymphatic vessels; HF, hair follicles; epi, epidermis; der, dermis. Histogram of the RT–qPCR analyses of the relative expression of Wls in the HFSC isolated from the K15CrePR+/T; WlsΔ/Δ mouse model and controls. n = 3–4 skin samples per mouse, n = 3–4 mice. Data represent the mean value ± SEM. ***P < 0.001 (Mann–Whitney U-test). Download figure Download PowerPoint Next, we explored if functional HFSC creates a niche promoting the continuous association of LV with the HF bulge. Wnt signaling features prominently in HFSC and regulates their properties and functional activity (Choi et al, 2013). Thus, we reduced the expression of Wnt ligands in HFSC by ablating the expression of Wls in the Keratin 15 (K15) HFSC compartment, using the K15CrePR+/T; WlsΔ/Δ conditional mouse model (Choi et al, 2013; Myung et al, 2013). Wntless (Wls) binds to Wnt ligands and controls their sorting and secretion (Carpenter et al, 2010). Under these conditions, HFSC remains quiescent, exhibiting a reduction in proliferation, but HF is largely maintained (Choi et al, 2013; Myung et al, 2013). Consistent with those findings, HF remained present in the skin (Fig 1G) despite the loss of Wls, as confirmed by RT–qPCR analyses of CD34+ α6 integrin+ FACS-isolated cells (Fig EV1B). LYVE1 immunofluorescence analyses revealed that the organized association of LV with HF was disrupted, and LV were found distributed parallel to the epidermis and distant from HF bulge areas (Fig 1G and H), reminiscent to the organization of LV in the ear skin (Fig EV1A). Overall, these results indicate that HFSC creates a niche for lymphatic endothelial cells, and sustain a polarized pattern of LV, which in turn, interconnect neighboring HF at the level of the HF bulge across the skin. Lymphatic capillaries start associating with HF during morphogenesis To gain further insight into the establishment of the LV–HF association, we next turned to HF embryogenesis. HFSC emerges in the early HF placode stage at E15.5, giving rise to the hair germ (E16.5), hair pegs (E17.5), and embryo Anagen HF until the first postnatal HF cycle. Whole mount immunofluorescence analyses of LYVE1 at E15.5–E17.5 embryo stages (Fig 2A–C) revealed that from E15.5, when HF placodes were already visible, nascent networks of anastomosed LV start to form below areas of HF growth. HF starts to develop via epithelial–mesenchymal inductive signals stemming from the dermal papilla (dp) (Sennett & Rendl, 2012); however, no apparent lymphatic association with the dp was observed at E15.5 or E16.5, and LV were rather aligned in parallel to the epidermis, representative of lymphatic collecting vessels (Fig 2A and B). Interestingly, at E17.5 upward grows of lymphatic capillaries started to branch out from collecting vessels toward the papillary dermis and distributed at HF sites (Fig 2C). Overall, these results indicate that the development of HF is coupled with the recruitment of lymphatic capillaries to HF sites, likely subsequent to HF specification. Figure 2. LV associate with HF during development A–C. Distribution of LV and HF at the mouse embryo stage E15.5 (A), E16.5 (B), and E17.5 (C). Maximum projection images of whole mount immunofluorescence analyses using LYVE1 (red) as lymphatic endothelial marker and counterstained with DAPI (blue). Red arrowheads denote HF placodes. n = 3–4 embryos. Scale bar, 50 μm. Download figure Download PowerPoint Lymphatic vessels interconnect triads of HF across the back skin To further analyze the organization of the lymphatic association with HF, we conducted whole mount immunofluorescence analyses of LYVE1 in adult skin sections (P70). This method allowed discerning an additional arrangement level, where individual LV–HF units associated further into triads (Fig 3A and Movie EV1). The patterned polygonal organization of lymphatic-HF domains across the skin was more evident in 3D projection planes (Fig 3B and Movie EV2). Lymphatic capillaries were found associated along the permanent portion of individual HF. At the level of the HF bulge, lymphatic capillaries radiated and converged interconnecting three HF units. These units presented a common extending lymphatic capillary from each HF triad (Fig 3B and Movie EV2). The mechanisms involved in the lymphatic-HF patterning are interesting questions for the future, but globally, these results uncharted the existence of coordinated lymphatic arrays surrounding and interconnecting triads of HF across the skin. Figure 3. LV association with HF during the postnatal HF cycle LV associated with individual HF further organize associating HF triads, which connect with other HF triads across the skin. P70, 70-day-old mice. n = 3–4 mice. White arrowheads denote capillaries stemming from HF triads. Scale bar, 100 μm. 3D reconstructions of whole skin mounts showing a triad of HF connected by LV. n = 3–4 mice. Scale bar, 50 μm. White arrowhead denotes an extending lymphatic capillary from a HF triad. Images of intravital microscopy analyses showing aligned HF rows in the back skin interconnected by LV in the Prox1CreERT2; Rosa-LSL-eYPF mice. Dotted boxes denote HF triads; white arrowheads denote capillaries stemming from HF triads interconnecting to other HF triads in adjacent rows. n = 3–4 mice. Scale bar, 50 μm. Download figure Download PowerPoint Lymphatic vessels continuously associate with the HF bulge during the HF cycle We next explored whether LV change their distribution to the HF bulge areas during the postnatal HF cycle. These analyses were performed in isolated back skin sections from matched skin areas at defined stages of the HF cycle, including the postnatal HF morphogenesis (postnatal days 5–16, P5–P16), and the first (P23–P45) and the second (P49–P85) HF cycle (Fig EV2; Muller-Rover et al, 2001). The latter exhibits a more extended Telogen that lasts for 3–4 weeks; therefore, to perform our comparative analyses we subdivided the second Telogen into early Telogen (Te, P49), mid-Telogen (Tm, P55), late Telogen (Tl, P69), and included an Anagen stage (AVI, P85; Muller-Rover et al, 2001). These phases corresponded to the refractory and competent Telogen phases, as previously documented (Plikus et al, 2008). Click here to expand this figure. Figure EV2. LV association with HF during the postnatal HF cycle Adult back skin sections from different postnatal (P) days immunostained for LYVE1 (red) and counterstained with DAPI (blue). n = 3–4 skin samples per mouse, n = 3–4 mice. Scale bar, 50 μm. epi, epidermis; der, dermis; LV, lymphatic vessels; HF, hair follicle. Histogram of the percentage of LYVE1-positive area in the HF permanent region at different postnatal days. Histogram of the percentage of LV length relative to the HF length in the back skin at different postnatal days. Histogram of the number of BrdU+ LYVE1+ cells/field in the back skin at different postnatal days. Histogram of the number of cleaved caspase-3+ LYVE1+ cells/field in the back skin at different postnatal days. Data information: The data shown in all histograms represent the mean value ± SEM. n = 3–4 skin samples per mouse, n = 3–4 mice. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001. (one-way ANOVA, Tukey's test). A, Anagen; C, Catagen; T, Telogen; Te, early Telogen; Tm, mid-Telogen; Tl, late Telogen. Download figure Download PowerPoint Interestingly, at all HF stages, LV remained distributed in a polarized manner, positioned at the anterior side of the permanent region of the HF opposite to the apm, interconnecting neighboring HF at the level of the HF bulge (Fig EV2A). The greater lymphatic density was localized at the HF permanent region, exhibiting a significantly higher area at postnatal HF morphogenesis stages (P5–P16; Fig EV2B). Consistent with this distribution, the relative LV/HF length increased from late Anagen stages to Telogen, when HF mostly consists of the permanent region (Fig EV2C). Conversely, this relative length was reduced during the transition from Telogen to Anagen (Fig EV2C). Except for the HF stages P5, P12, and P16, no changes in LV proliferation were observed (Fig EV2D), suggesting that LV were still growing and reorganizing to developing HF. Also, no changes in LV cell death were observed during the HF cycle (Fig EV2E). These studies exposed that LV establish a continuous connection between neighboring HF at the level of the HF bulge across the skin throughout all phases of the HF cycle. Lymphatic vessels dynamically flow across triads of HF in the back skin To further pursue the existence of dynamic lymphatic flow between neighboring HF triads across the skin, we analyzed the lymphatic network by intravital microscopy, using a Prox1-CreERT2; ROSA26-LSL-eYFP reporter mouse. This transgenic mouse line expresses the tamoxifen-inducible Cre recombinase (CreER) under the control of the Prox1 gene promoter (Bazigou et al, 2011), under a Rosa26-LSL-eYPP background. Prox1 is essential for LV development, and it is expressed throughout life, providing lymphatic identity (Wigle & Oliver, 1999; Hong et al, 2002). These powerful tools allowed us to explore the dynamic association of lymphatic capillary networks with HF in living skin tissue. Our intravital microscopy analyses fully evidenced the continuum-patterned organization of lymphatic networks around HF in the back skin (Fig 3C). These analyses further allowed the visualization of triads of HF interconnected by lymphatic capillaries, aligned in parallel rows with an anterior to posterior disposition. Strikingly, the HF triads in each row interconnected with neighboring parallel rows, mainly through a LV stemming from one HF triad unit (Fig 3C), consistent with our prior observations (Fig 3B and Movie EV2). We next assessed whether LV dynamically streamed into the continuum-patterned lymphatic networks around HF in the back skin. To this end, TRITC–Dextran was administered in the mouse back skin followed by intravital microscopy. These results allowed the visualization of a continuous lymphatic flow interconnecting adjacent HF rows (Movie EV3). These results exposed the existence of a dynamic HF communication through lymphatic vascularization, which potentially facilitates the spreading of signaling waves and immune cell trafficking across HF in the back skin. Lymphatic endothelial cells transiently increase their caliber at the onset of HF stem cell activation Our previous findings raised the possibility that LV undergo dynamic flow changes at different stages of the HF cycle. We investigated this aspect by measuring the LV caliber in back skin sections during the first and the second HF cycle, conducting LYVE1 immunofluorescence analyses. Intriguingly, the LV caliber was more pronounced at the onset of the Telogen to Anagen transition (Fig 4A and B), without exhibiting signs of proliferation or cell death (Fig EV2D and E). We next inspected more closely the LV morphology and observed that at late stages of Telogen, LV appeared more fenestrated displaying membrane protrusions compared to the more continuous and tight capillaries observed during Anagen. This structural variation, exhibiting a wide and irregular lumen, presumably accounts for regional differences in capillary permeability (O'Driscoll, 1992) influencing vascular exchange and increase tissue drainage capacity (Aebischer et al, 2014; Betterman & Harvey, 2016). Figure 4. Dynamic reorganization of LV during the HF cycle A. Adult back skin sections from different postnatal (P) days immunostained for LYVE1 (red) and counterstained with DAPI (blue). n = 3–4 skin samples per mouse, n = 3–4 mice. Scale bar, 10 μm. White arrowheads denote LV membrane protrusion and fenestrated areas. LV, lymphatic vessels; HF, hair follicle. B. Histogram of the caliber (μm) of LV at different postnatal days. n = 3–4 skin samples per mouse, n = 3–4 mice. A, Anagen; C, Catagen; T, Telogen; Te, early Telogen; Tm, mid-Telogen; Tl, late Telogen. The data shown represent the mean value ± SEM. *P < 0.05; **P < 0.01; ***P < 0.001 (one-way ANOVA, Tukey's test). C. H&E staining of back skin sections from mice treated intradermally with Control or Clodronate liposomes. n = 3–4 skin samples per mouse, n = 3–4 mice. Scale bar, 200 μm. epi, epidermis; der, dermis. D, E. LYVE1 immunofluorescence (red) counterstained with DAPI (blue) (D) and histogram of the caliber (μm) of LV (E) in back skin sections from mice treated intradermally with Control or Clodronate liposomes. n = 3–4 skin samples per mouse, n = 3–4 mice. Scale bar, 100 μm. epi, epidermis; der, dermis. The data shown represent the mean value ± SEM. ****P < 0.0001 (Mann–Whitney U-test). F. Back skin sections of Controls and K14ΔNβ-cateninER+/T mice immunostained for LYVE1 (red) and counterstained with DAPI (blue). n = 3–4 skin samples per mouse, n = 3–4 mice. Scale bar, 50 μm. epi, epidermis; der, dermis. G. Histogram of the LV caliber in skin sections from Controls and K14ΔNβ-cateninER+/T mice. n = 3–4 skin samples per mouse, n = 3–4 mice. Data represent the mean value ± SEM. ****P < 0.0001 (Mann–Whitney U-test). H. Histogram of the percentage of HF in Telogen and Anagen present in the skin of control mice or mice treated with Cilostazol. n = 3–4 skin samples per mouse, n = 3–4 mice. Data represent the mean value ± SEM. *P < 0.05; **P < 0.01 (Mann–Whitney U-test). Download figure Download PowerPoint We next explored whether the induction of HF growth induces the transitory increase in the caliber of LV. To this end, to avoid severing LV and the induction of an inflammatory response, we did not conduct hair plucking experiments to synchronize HF, but rather stimulated the Telogen to Anagen transition by reducing the number of perifollicular macrophages at early Telogen with Clodronate liposomes (Fig 4C–E), as previously described (Castellana et al, 2014). Under these conditions, LV surrounding the precocious Anagen HF displayed an increase in their caliber compared to controls (Fig 4D and E). We further analyzed the connection between HFSC cell proliferation and the expansion of LV caliber in K14Cre+/T, ΔNβ-cateninlox/lox mouse skin samples. This model i
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