Research Article

Dermal sheath contraction powers stem cell niche relocation during hair cycle regression

See allHide authors and affiliations

Science  10 Jan 2020:
Vol. 367, Issue 6474, pp. 161-166
DOI: 10.1126/science.aax9131

Niche relocated by muscle contraction

Regulation of adult stem cells by their microenvironment, or niche, is essential for tissue homeostasis and for regeneration after injury and during aging. Normal regression of hair follicles during the hair cycle poses a particular challenge for maintaining a functional proximity of stem cells to their niche, especially the specialized mesenchymal cells of the dermal papilla. Using mice as a model organism, Heitman et al. demonstrate that the follicle dermal sheath is an active smooth muscle that drives tissue remodeling through coordinated cell contraction, enabling renewed contact between the dermal papilla and hair follicle stem cells during hair follicle regression. This biomechanical mechanism of niche relocation may be utilized in other stem cell niche systems.

Science, this issue p. 161


Tissue homeostasis requires the balance of growth by cell production and regression through cell loss. In the hair cycle, during follicle regression, the niche traverses the skin through an unknown mechanism to reach the stem cell reservoir and trigger new growth. Here, we identify the dermal sheath that lines the follicle as the key driver of tissue regression and niche relocation through the smooth muscle contractile machinery that generates centripetal constriction force. We reveal that the calcium-calmodulin–myosin light chain kinase pathway controls sheath contraction. When this pathway is blocked, sheath contraction is inhibited, impeding follicle regression and niche relocation. Thus, our study identifies the dermal sheath as smooth muscle that drives follicle regression for reuniting niche and stem cells in order to regenerate tissue structure during homeostasis.

Multipotent progenitors produce new cells to replace those that have been lost by injury or by natural turnover and shedding in rapidly renewing tissues such as the bone marrow and hair follicle (1). However, progenitors have limited renewal capacity and must be replenished from the stem cell reservoir, which is coordinated with specialized niche cells (24). During hair growth, the specialized dermal papilla (DP) cluster at the base of follicle bulbs secretes niche signals to orchestrate matrix progenitor proliferation, migration, and differentiation (5, 6) (fig. S1A). To reboot the progenitor pool throughout life, repeated cycles of follicle regression (catagen), rest (telogen), and regrowth (anagen) (7) (fig. S1B) derive new cells from the stem cell reservoir located in the bulge and germ of the upper follicle (812). At the beginning of regression, matrix progenitors stop proliferating and terminally differentiate (fig. S1B). Throughout regression, then, nearly all outer root sheath (ORS) progenitors along the entire length of catagen follicles undergo apoptotic pruning (13) (Fig. 1A and fig. S1B) that is driven by extrinsic signals (14, 15). Only the ORS progenitors located closely below the bulge stem cells survive and retain multipotency to give rise to a new bulge and germ (16, 17). This new stem cell reservoir then becomes reactivated by niche signals from the intact, surviving DP after it has relocated from the follicle base to the new bulge and germ in order to regenerate the follicle of the next growth cycle (1820) (fig. S1B).

Fig. 1 The dermal sheath is required for hair follicle regression.

(A) Schematic of catagen regression during the hair cycle. Hair shaft, inner root sheath, and matrix progenitors are eliminated by terminal differentiation and extrusion from the skin. The majority of ORS progenitors (blue) are eliminated by apoptosis. It is unknown how the surviving DP niche relocates to the bulge and germ stem cell reservoir to activate new hair growth after a period of rest. (B) Immunofluorescence (IF) for ACAN, secreted by ITGA8+ DS cells. The right image is a magnification of the region enclosed by the white dashed line in the left image. DAPI, 4′,6-diamidino-2-phenylindole. (C) Schematic of cytotoxic DS ablation during regression. tam, tamoxifen; P, postnatal day. (D) IF for αSMA in control (R26LSL-DTA) and DS-ablated (AcanCreER;R26LSL-DTA) back skin. The white dashed lines indicate the location of the basement membrane between ORS and DS. (E) Whole-mount IF for ORS marker K14 in P20 back skins (viewed from the dermis side, anterior is to the left). Control follicles are in telogen resting phase. Note the elongated follicles after DS ablation that are stalled in the regression phase. The white dashed box indicates the region shown in (G). (F) Quantification of the percentage of stalled follicles at P20 (n = 698 follicles in control and 895 follicles in DS-ablated skin in five mice). **P = 0.003, by unpaired two-tailed Student’s t test. (G) Magnification of the region enclosed by the white dashed line in (D) also stained with DAPI showing that the hair shaft and DP remain at the bulb tip of stalled follicles. (H) Quantification of follicle lengths [n = 11 for P13 control (Co.), n =14 for P13 ablated (Abl.), n = 80 for P20 control, and n = 27 for P20 stalled; 11 mice]. ****P < 10−4, by unpaired two-tailed Student’s t test. (I) Stalled follicles have no DS (αSMA) but retain intact DP (LEF1+). K14+ ORS progenitor and K6+ companion (Cp) and medulla (Me) layers are present and lack apoptosis (activated CASP3) or proliferation (Ki67) markers. The white dashed lines indicate the location of the basement membrane. Scale bars are 50 μm [(B) and (E)] and 10 μm [(D) and (I)].

In relocating to the stem cell reservoir, the DP trails the regressing epithelial strand of dying ORS progenitors and travels more than 90% of the hair follicle length (fig. S2) toward the bulge and germ stem cell reservoir. Such drastic tissue remodeling poses a considerable challenge for stem cell regulation in establishing the required proximity to its niche for receiving activating signals. Indeed, detachment of the DP leads to stem cell dysfunction and hair loss (21, 22). Two hypotheses for DP movement have been proposed for many years (23): (i) an “apoptotic force” generated by dying cells in the shrinking follicle and (ii) a “contraction force” by follicle-encapsulating dermal sheath (DS) cells (Fig. 1A and fig. S1B) based on alpha smooth muscle actin (αSMA) expression identified nearly three decades ago (24).

Dermal sheath is essential for reuniting stem cells and niche

We set out to address whether the DS is important for niche relocation during follicle regression. We first established DS-specific targeting using cartilage proteoglycan Aggrecan (Acan) as a genetic driver. ACAN protein is found in skin only in the follicle basement membrane that separates the DS from epithelium (25) (Fig. 1B). Using AcanCreER mice crossed with the R26LSL-tdTomato reporter (AcantdT) enabled inducible targeting during catagen regression, restricted within skin to the DS (fig. S3). To test its functional role, we selectively ablated the DS at early catagen using inducible diphtheria toxin driver R26LSL-DTA (AcanDTA) (Fig. 1C). In follicles with effective ablation, only DS cell remnants were left at the beginning of regression (Fig. 1D). By subsequent telogen rest, follicles in control skin were fully regressed (Fig. 1E) and follicles with inefficient DS ablation equally regressed (fig. S4), but many follicles with fully ablated DS (fig. S4) remained aberrantly long (Fig. 1, E and F), suggesting a threshold requirement of minimal remaining DS for regression. In stalled follicles, hair shafts extended down to the follicle base and DPs remained deep in the dermis (Fig. 1G). Stalled follicle lengths ranged from partially regressed to completely stalled deep within the dermis (Fig. 1H). This indicates that the DS is required for progression of follicle regression during catagen and for niche relocation to the stem cell reservoir in the upper follicle.

We next analyzed the stalled follicle phenotype more closely with lineage-specific markers (figs. S1A and S5A) to determine the degree of catagen dysfunction in nonregressing hair follicles lacking DS. Immunofluorescence confirmed the absence of the DS and widespread ORS cells around shafts of stalled follicles (Fig. 1I). Rounded DP cell clusters at the bulb end were no longer engulfed by proliferative matrix progenitors (Fig. 1I). Likewise, hair shaft and inner root sheath precursors were also absent (fig. S5B), because differentiation during anagen growth (fig. S5A) had ended and catagen regression was initiated. Apoptosis was also undetectable in DS-ablated follicles (Fig. 1I), suggesting that catagen did not proceed beyond the early steps and that normal catagen cell death (fig. S5C) was absent. Between hair shafts and ORS was a single differentiated layer (Fig. 1I), reminiscent of the companion layer in growing hair follicles (fig. S5A) and of inner bulge cells at telogen (fig. S5D). Together, the phenotypic analyses indicated that long, nonregressed follicles were not caused by continued matrix progenitor proliferation or erroneous differentiation but resulted from a failure of hair shaft and DP movement toward the skin surface and concomitant absence of ORS progenitor apoptosis. Thus, the DS may exert a physical force fundamental for the upward travel of hair shaft and DP niche.

Dermal sheath expresses the molecular machinery of smooth muscles

To explore if the DS expresses the contractile molecular machinery of smooth muscles that could execute follicle regression and niche relocation, we isolated DS cells for transcriptome analysis, in comparison to DP and dermal fibroblasts (DFs). We flow-sorted DS, DP, and DF populations from cells that were isolated from digested back skins of Sox2GFP;Lef1-RFP reporter mice (6, 26) and were also stained for mesenchymal receptor platelet-derived growth factor receptor A (PDGFRA) (Fig. 2, A and B, and fig. S6, A and B). DS cells expressed Sox2GFP and PDGFRA, but not red fluorescent protein (RFP), and could be cleanly distinguished from DP and DFs (27) (Fig. 2, A and B). Verification of known marker expression for DS, DP, and DFs demonstrated their identity and purity (fig. S6C). Analyzing the RNA sequencing–generated transcriptomes of DS, DP, and DFs and of ORS, matrix, and melanocytes (27) by hierarchical clustering and principal components established their close lineage relationship (fig. S7, A and B). Comparative analysis of gene expression then revealed a DS molecular signature of 483 enriched genes reflecting their specialized functions (Fig. 2C, fig. S7C, and table S1). Gene ontology analysis of the DS signature expectedly yielded extracellular matrix organization categories (Fig. 2D), because the DS is closely associated with the basement membrane that separates the mesenchyme from follicle epithelium. The DS signature was also enriched for genes involved in “muscle filament sliding” and “smooth muscle contraction” (Fig. 2D). Gene set enrichment analysis for “regulation and process of smooth muscle contraction” (table S2) showed significant enrichment in the DS (Fig. 2E), suggesting smooth muscle identity and function.

Fig. 2 The dermal sheath expresses the molecular machinery of smooth muscles.

(A and B) Labeling of DS and DP (A) and flow cell sorting (B) from Sox2GFP;Lef1-RFP P5 back skin after IF for PDGFRA. DFs were sorted for comparison. (C) Venn diagram of gene signatures. (D) Gene ontology analysis of DS signature. (E) Gene set enrichment analysis (GSEA) for genes involved in “smooth muscle contraction and regulation” are highly enriched in DS. NES, normalized enrichment score; FDR, false discovery rate. (F) Schematic of Ca2+-dependent smooth muscle contraction pathway. (G) Heatmap of smooth muscle contraction gene expression. Ca2+ contraction pathway and pan–smooth muscle genes (asterisks) are highly enriched in DS. (H) 3D IF for αSMA fibers arranged in a concentric ring–like network wrapping around the follicle. The inset is a magnification of the area enclosed by the white dashed line. (I) IF of smooth muscle contraction components in DS. Scale bars are 50 μm.

Several core components of the Ca2+-dependent smooth muscle contraction pathway (Fig. 2F) were highly enriched in the DS, including Calm1 (calmodulin, CaM), Mylk (myosin light chain kinase, MLCK), Myh11 (myosin heavy chain 11, MYH11), Myl9 (myosin light chain 9, MYL9), and Acta2 (αSMA) (Fig. 2G). Several pan–smooth muscle genes (28), such as Cald1 (caldesmon 1), Smtn (smoothelin), and Tagln (transgelin, SM22), were also part of the DS signature (Fig. 2G and fig. S7, D and E). Among these, Cald1 and Smtn are not expressed in contractile myofibroblasts (29), indicating that the DS expresses genes of bona fide smooth muscles. During smooth muscle contraction, mechanical forces are generated through actomyosin cross-bridges and adenosine triphosphate (ATP)–powered myosin ratcheting action (30) (Fig. 2F). Three-dimensional (3D) immunofluorescence revealed that the DS forms a network of αSMA stress fibers that wrap the follicle in concentric rings (Fig. 2H), suggesting that the potential actomyosin forces would be directed toward the center of the longitudinal axis in centripetal fashion. Actomyosin cross-bridge formation is promoted when CaM-activated MLCK phosphorylates regulatory myosin light chains (e.g., MYL9; Fig. 2, F, G, and I) that associate with myosin heavy chain molecular motors (e.g., MYH10; Fig. 2, F, G, and I). Expression of phosphorylated MYL9 (pMYL9) confirms the active state of myosin in the entire length of DS (Fig. 2I), which we observed throughout catagen (fig. S8), and supports DS functional contractile activity. Finally, several smooth muscle proteins were expressed in human scalp hair follicles, suggesting conservation of smooth muscle components in the DS between mice and humans (fig. S9). Overall, these results demonstrate that the DS harbors the contractile apparatus and its regulatory elements, long hypothesized by the presence of αSMA (24).

Dermal sheath contraction is required for follicle regression

We next explored whether DS cells can functionally contract in a smooth muscle–like fashion through Ca2+-dependent MLCK activation. Isolated, short-term cultured DS cells were membrane depolarized by an extracellular K+ spike to activate voltage-gated Ca2+ channels in the presence of Fluo8 fluorescent Ca2+ indicator (fig. S10A). The switch from Na+ to high-K+ media led to effective Ca2+ influx within 1.5 min. Tracing the cell surface area of AcantdT-marked DS cells cultured on a soft substrate demonstrated functional contraction in high-K+ conditions in vitro (fig. S10B). Next, we tested whether the DS can functionally contract and compress microdissected intact hair follicles (Fig. 3A). Live imaging of freshly isolated follicles showed significant reduction of follicle widths after 3 min in high K+, consistent with contraction of concentric αSMA-myosin rings (Fig. 3, B and D, and movie S1), which was effectively blocked by preincubation with the MLCK-specific inhibitor ML7 (Fig. 3, C and D, and movie S2). These data demonstrate that the DS can functionally contract by activation of voltage-gated Ca2+ channels in the CaM → MLCK → MYL-MYH-αSMA pathway.

Fig. 3 The dermal sheath functionally contracts and is required for regression in vivo.

(A) Schematic of live-imaging microdissected follicles preincubated with or without MLCK inhibitor ML7 and after high K+ depolarization. (B and C) Still images from bright-field movie at start (black) and end (pink) of high K+ incubation in the absence of inhibitor (B) and in the presence of ML7 (C). Overlays highlight reduction of follicle width, blocked by ML7. (D) Quantification of follicle widths during live imaging. n = 7 follicles for ML7 and no inhibitor preincubation. Data points are mean ± SD. **P < 0.01, by unpaired two-tailed Student’s t test. (E to G) Topical inhibition of MLCK by ML7 blocks hair follicle regression in vivo. Schematic of ML7 or vehicle (DMSO, dimethyl sulfoxide) application during catagen (E). Whole-mount IF of P20 back skins shows normal regression of follicles into telogen rest in control but stalled follicles in contraction-inhibited ML7-treated regions (F). Quantification of the percentage of stalled follicles (n = 1071 control, and n = 1019 ML7-treated; 10 mice) (G). Data bars are mean ± SD. **P = 0.001, by unpaired two-tailed Student’s t test. (H) IF for LEF1, Ki67, αSMA, and K14. Stalled follicles have intact DS (αSMA) and DP (LEF1) that are no longer engulfed. Epithelial cells (K14) of stalled follicles are not proliferative (Ki67). Scale bars are 50 μm [(B), (C), and (F)] and 10 μm (H).

To determine whether smooth muscle–like contraction is a main function of the DS to propagate hair follicle regression in vivo, we blocked the MLCK pathway by topical ML7 application on the back skin of mice throughout catagen regression (Fig. 3E). In the back skins of mice receiving vehicle control, follicles regressed normally into telogen with the DP relocated next to the stem cell reservoir (Fig. 3F and fig. S10C). Blocking smooth muscle contraction with ML7, however, resulted in a notable failure of regression with long stalled follicles stranded deep in the dermis in the center of the treated skin (Fig. 3, F and G), reminiscent of the DS-ablated phenotype (Fig. 1E). The DP here also remains at the tip of stalled follicles (Fig. 3H), but in contrast to ablated follicles, the intact DS surrounds follicles including DP. Taken together, these data demonstrate that DS contraction is functionally required for follicle regression and DP niche relocation to the stem cells.

Visualizing regression movements by intravital imaging

By utilizing intravital two-photon microscopy of unperturbed skin in live mice (15, 18) (fig. S11A), we investigated where the DS contracts and by which force mechanism the DS drives niche relocation. For this, we first established triple-color fluorescent reporters to visualize both the DS and neighboring progenitors (Fig. 4A). AcantdT labels the flat DS cell cytoplasm in red, and Tbx18H2BGFP (31) labels DS and DP nuclei in bright and dim green. To additionally label all nuclei of neighboring progenitors bright blue, we generated a K14-H2BCer transgenic line (Fig. 4A and fig. S12, A and B). The triple-color fluorescent reporter combination (fig. S12C) enabled simultaneous live imaging of DS, DP, and follicle epithelium at a single excitation wavelength (fig. S11, B and C) and thus fine-mapping of their absolute and relative movement over time.

Fig. 4 Dermal sheath contraction is required for hair shaft and niche relocation.

(A) Triple-fluorescent reporter follicles for intravital 3D time-lapse imaging of catagen regression in live mice (5 hours). AcantdT and Tbx18H2BGFP mark DS cytoplasm and nuclei, respectively. K14-H2BCer highlights all epithelial nuclei. The DP was recognized by low-level Tbx18H2BGFP expression and surrounding epithelial and DS cells. (B) Upward movement of hair shaft and DP during regression. Tracking of individual ORS, shaft, and DS cells and of upper and lower bounds of DP during 5-hour imaging. The white arrows indicate distance traveled of shaft and DP. (C) Quantification of live cell tracking relative to ORS movement (7.5-min intervals). Shaft and DP move upward relative to ORS and DS. Solid lines are averages; shaded areas are SD. n = 13 hair shaft, n = 8 DP, n = 26 ORS, and n = 17 DS measurements (seven follicles, three imaging sessions). (D) In vivo contraction blocking during intravital imaging in K14-H2BCer mice. Inverted grayscale still images at beginning and end of vehicle control followed by ML7 application. Quantification of live tracking of shaft cells (black) relative to ORS (blue). n = 9 follicles from two independent imaging sessions. **P < 0.01, and ***P < 0.001, by paired two-tailed Student’s t test. (E) Schematic of two historically hypothesized mechanisms for DP niche relocation during regression. (F and G) Fluorescence images from time-lapse movie (F) and quantification (G) of the length of regressing epithelial strand. Strand lengths remain unchanged and stable (n = 10 follicles). P = 0.572, by unpaired two-tailed Student’s t test (0 versus 5 hours). (H) DS cross-sectional diameter at DP bottom (n = 4 follicles). P = 0.994, by unpaired two-tailed Student’s t test (0 versus 5 hours). (I) Quantification of DS diameter that remains unchanged over time. Data bars are mean ± SD. Scale bars are 10 μm.

Cell tracking during a 5-hour span of catagen regression demonstrated upward movement of the hair shaft and DP, as expected (Fig. 4B, fig. S13A, and movie S3). By contrast, the DS and ORS showed minimal movement in the long axis (Fig. 4C and fig. S13, B and C), indicating that hair shaft upward movement occurs at the interface between ORS progenitors and shaft, as previously observed (15). The lack of relative movement between DS and ORS suggested strong connections through the separating basement membrane. We next visualized hair shaft regression movements directly after contraction blocking in vivo (Fig. 4D). Live imaging the same follicle before and after topical ML7 application demonstrated that blocking smooth muscle contraction effectively abrogated upward movement of hair shafts. Together, our labeling setup and live-imaging time frame effectively captured the movements of key components during catagen regression and confirmed the requirement of smooth muscle contraction.

We then asked by which mechanism the niche becomes relocated during regression to reach its final position next to the stem cell reservoir and tested two long-standing hypotheses (24) (Fig. 4E): (i) an apoptotic force from the epithelial strand pulls the DP (23) and (ii) the DS contracts to push the DP and/or hair shaft upward. To investigate the first hypothesis, we measured the length of the epithelial strand during the 5-hour imaging window. If shrinking due to dying cells would generate force for pulling the DP, we would expect shortening of the strand correlating to DP movement (Fig. 4E). However, we observed that individual epithelial strand lengths remained constant throughout (Fig. 4, F and G) despite steady upward DP movement, consistent with similar rates of shaft and DP upward movement (Fig. 4C). Although forces from individual migrating epithelial strand cells could contribute to upward DP movement, our findings suggest that apoptotic force is not a major driver of regression.

To test the hypothesis that a DS-generated contractile force pushes the DP from underneath, we next investigated potential DS contraction movement below the DP. Coinciding with DP upward movement, we would expect DS centripetal movement underneath the DP. Therefore, we measured the cross-sectional diameter of the DS over time, which, however, remained unchanged (Fig. 4, H and I), suggesting that direct DS contraction under the DP likely does not account for its relocation.

Contraction at the follicle bottleneck pulls the niche via the epithelial strand

We then considered a variation of the contraction hypothesis, in which DS contraction generates constriction forces that push the shaft upward, which in turn would pull both the epithelial strand and the DP upward (Fig. 5A). The observation of a constant epithelial strand length over time (Fig. 4, F and G) supports this alternative hypothesis in which the strand acts as a stable tether between the shaft and DP (Fig. 4F). Indeed, centripetal contraction movement of DS cells right at the border that forms a “bottleneck” between the shaft-containing club hair and the narrower epithelial strand (Fig. 5, B and C) indicates that DS contraction pushes the shaft upward (Fig. 5B and movie S3). After measuring the cross-sectional diameter of the follicle-wrapping DS cell ring at this bottleneck, we found a significant decrease over time coinciding with shaft upward movement (Fig. 5, D and E). These data reveal that the DS moves centripetally at the bottleneck toward the center, suggesting that it contracts to generate the constriction force necessary for pushing the hair shaft upward—akin to the squeezing motion of a toothpaste tube—which then pulls the DP by its connection to the hair shaft via the epithelial strand (Fig. 5F).

Fig. 5 Dermal sheath contraction pushes the hair shaft and indirectly pulls the niche.

(A) Schematic of a third, alternative hypothesis for DP niche relocation by DS contraction at the bottleneck between the shaft-containing club hair and narrower regressing epithelial strand. (B) Live cell tracking of DS centripetal constriction movement at club hair–epithelial strand bottleneck and of hair shaft upward movement. White arrows are starting and ending positions of 5-hour tracking (seven follicles, three imaging sessions). (C) High magnification of fluorescence time-lapse images at club hair–epithelial strand bottleneck. (D) DS cross-sectional diameter at club hair–epithelial strand bottleneck. The diameter of the follicle-wrapping DS cell ring decreased over time. (E) Quantification of DS diameter at bottleneck decreases over time (n = 3 follicles). *P = 0.021, **P = 0.009, and P = 0.010, by unpaired two-tailed Student’s t test (0 versus 5 hours). Data bars are mean ± SD. (F) Model for DP niche relocation during regression. DS contraction forces centripetally constrict the follicle at the bottleneck to move the hair shaft upward, which pulls the DP upward via the epithelial strand. Scale bars are 10 μm.

Concluding remarks

Here, we answer the long-standing question of how the niche is relocated nearly the entire length of the follicle to reach its essential stem cell–adjacent position. Through intravital imaging, contraction assays in isolated cells and intact follicles ex vivo, and in vivo inhibition of contraction, we demonstrate that DS is a smooth muscle that contracts to power the key follicle regression movements during the catagen phase of the hair cycle. We directly tested the hypotheses that proposed apoptotic or contraction forces propel catagen progression, based on observations possible at the time, and consolidated them into a model in which the niche relocates to the stem cells through a series of force relays (Fig. 5F). As an adaptation of the original contraction hypothesis, the DS contracts throughout catagen at the club hair–epithelial strand bottleneck, where, because of its angle, the centripetal constricting force gets redirected to an upward pushing force on the shaft. This force is then relayed through the tether-like epithelial strand pulling the DP. At the end stage of regression, when the hair shaft has reached its final position and the DS trails below the DP as a hollow sleeve, it is possible that forces generated by the space reduction of apoptosing cells then pull in the niche next to the stem cell reservoir before launching regrowth in the next cycle that regenerates the follicle for new hair shaft production.

Intimate cross-talk between stem cells and their niches is vital for proper stem cell maintenance and cell fate decisions. Therefore, it is not surprising that niches are found in anatomically distinct locations for hematopoietic stem cells in the bone marrow and intestinal stem cells at the crypt base. Providing close proximities for paracrine signaling and insulation from outside influences, most stem cell niche systems under homeostasis remain structurally stable, but after injury to the niche, reestablishment is vital for the restoration of long-term function (32, 33). Although hematopoietic stem cells can mobilize and home to their niche, such as during transplantation (34), the more restricted freedom of movement of epithelial stem cells may limit their homing potential. Here, we discovered another smooth muscle function—relocating a niche to its stem cell reservoir—to add to the vast array of diverse roles of smooth muscles throughout the body. This example highlights the evolutionary advantage of repurposing preexisting functionality rather than inventing new systems in the wake of new adaptive challenges. Further study may identify more instances of the function of smooth muscle contraction in regulating stem cell–niche signaling range in homeostasis, which could be lost or exploited in disease.

Supplementary Materials

Materials and Methods

Figs. S1 to S13

Tables S1 to S3

References (3544)

Movies S1 to S3

References and Notes

Acknowledgments: We thank V. Horsley, T. Tumbar, R. Paus, M. Rangl, and R. Krauss for invaluable discussions and comments on the manuscript. We are especially grateful to C. Jahoda for helpful discussions and launching dermal sheath research decades ago. Many thanks to D. Dubin and H. Khorasani for providing human scalp skin samples and to the personnel at ISMMS Flow Cytometry, Microscopy, Genomics, and Mouse Genetics CoREs for technical assistance. Funding: The ISMMS Microscopy CoRE was supported by NIH Shared Instrumentation grant IS10RR026639. N.H. and R.S. were supported by training grant T32GM007280 from NIH/NIGMS. N.H. was also supported by T32HHD075735 from NIH/NIDCR and F30AR070639 from NIH/NIAMS. D.S. and N.S. were supported by a fellowship of the Training Program in Stem Cell Research from the New York State Department of Health (NYSTEM-C32561GG). K.-W.M. was supported by The Science Appearance Career Development Award fellowship from the Dermatology Foundation. A.M. was supported by grants from NIH/OD (U54HL127624 and U24CA224260). P.R. was supported by an NIH/NEI grant (R01EY030599). M.R. was supported by grants from NIH/NIAMS (R01AR071047 and R01AR063151) and the New York State Department of Health (NYSTEM-C029574 and NYSTEM-C32561GG) and by a fellowship from the Irma T. Hirschl Trust. Author contributions: N.H., R.S., and M.R. designed the experiments and overall study. N.H. and M.R. wrote the manuscript. N.H., D.S., R.S., K.-W.M., N.S., P.M., and L.G. performed the experiments. Z.W. and A.M. assisted with transcriptome analysis. P.R. assisted with two-photon imaging. All authors discussed the results and participated in the manuscript preparation and editing. M.R. supervised the study. Competing interests: N.H., R.S., and M.R. are inventors on a confidential provisional patent application that covers the subject matter of this article. The authors declare no other competing interests. Data and materials availability: All data presented in this study are available in the main text or the supplementary materials. Raw and analyzed RNA-sequencing data generated during this study are available in the Gene Expression Omnibus (GEO) repository under accession GSE136996.

Stay Connected to Science

Navigate This Article