Hair follicle stem cell cultures reveal self‐organizing plasticity of stem cells and their progeny
2016; Springer Nature; Volume: 36; Issue: 2 Linguagem: Inglês
10.15252/embj.201694902
ISSN1460-2075
AutoresCarlos Andrés Chacón‐Martínez, Markus Klose, Catherin Niemann, Ingmar Glauche, Sara A. Wickström,
Tópico(s)Skin and Cellular Biology Research
ResumoArticle9 December 2016free access Transparent process Hair follicle stem cell cultures reveal self-organizing plasticity of stem cells and their progeny Carlos Andrés Chacón-Martínez Carlos Andrés Chacón-Martínez Paul Gerson Unna Group "Skin Homeostasis and Ageing", Max Planck Institute for Biology of Ageing, Cologne, Germany Search for more papers by this author Markus Klose Markus Klose Institute for Medical Informatics and Biometry, Carl Gustav Carus Faculty of Medicine, Technische Universität Dresden, Dresden, Germany Search for more papers by this author Catherin Niemann Catherin Niemann Institute for Biochemistry II, Medical Faculty, University of Cologne, Cologne, Germany Center for Molecular Medicine Cologne, University of Cologne, Cologne, Germany Search for more papers by this author Ingmar Glauche Ingmar Glauche Institute for Medical Informatics and Biometry, Carl Gustav Carus Faculty of Medicine, Technische Universität Dresden, Dresden, Germany Search for more papers by this author Sara A Wickström Corresponding Author Sara A Wickström [email protected] orcid.org/0000-0001-6383-6292 Paul Gerson Unna Group "Skin Homeostasis and Ageing", Max Planck Institute for Biology of Ageing, Cologne, Germany Cologne Excellence Cluster on Cellular Stress Responses in Aging-Associated Diseases (CECAD), University of Cologne, Cologne, Germany Search for more papers by this author Carlos Andrés Chacón-Martínez Carlos Andrés Chacón-Martínez Paul Gerson Unna Group "Skin Homeostasis and Ageing", Max Planck Institute for Biology of Ageing, Cologne, Germany Search for more papers by this author Markus Klose Markus Klose Institute for Medical Informatics and Biometry, Carl Gustav Carus Faculty of Medicine, Technische Universität Dresden, Dresden, Germany Search for more papers by this author Catherin Niemann Catherin Niemann Institute for Biochemistry II, Medical Faculty, University of Cologne, Cologne, Germany Center for Molecular Medicine Cologne, University of Cologne, Cologne, Germany Search for more papers by this author Ingmar Glauche Ingmar Glauche Institute for Medical Informatics and Biometry, Carl Gustav Carus Faculty of Medicine, Technische Universität Dresden, Dresden, Germany Search for more papers by this author Sara A Wickström Corresponding Author Sara A Wickström [email protected] orcid.org/0000-0001-6383-6292 Paul Gerson Unna Group "Skin Homeostasis and Ageing", Max Planck Institute for Biology of Ageing, Cologne, Germany Cologne Excellence Cluster on Cellular Stress Responses in Aging-Associated Diseases (CECAD), University of Cologne, Cologne, Germany Search for more papers by this author Author Information Carlos Andrés Chacón-Martínez1, Markus Klose2, Catherin Niemann3,4, Ingmar Glauche2 and Sara A Wickström *,1,5 1Paul Gerson Unna Group "Skin Homeostasis and Ageing", Max Planck Institute for Biology of Ageing, Cologne, Germany 2Institute for Medical Informatics and Biometry, Carl Gustav Carus Faculty of Medicine, Technische Universität Dresden, Dresden, Germany 3Institute for Biochemistry II, Medical Faculty, University of Cologne, Cologne, Germany 4Center for Molecular Medicine Cologne, University of Cologne, Cologne, Germany 5Cologne Excellence Cluster on Cellular Stress Responses in Aging-Associated Diseases (CECAD), University of Cologne, Cologne, Germany *Corresponding author. Tel: +49 221 379 70 770; E-mail: [email protected] The EMBO Journal (2017)36:151-164https://doi.org/10.15252/embj.201694902 See also: A Sánchez-Danés & C Blanpain (January 2017) 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 Understanding how complex tissues are formed, maintained, and regenerated through local growth, differentiation, and remodeling requires knowledge on how single-cell behaviors are coordinated on the population level. The self-renewing hair follicle, maintained by a distinct stem cell population, represents an excellent paradigm to address this question. A major obstacle in mechanistic understanding of hair follicle stem cell (HFSC) regulation has been the lack of a culture system that recapitulates HFSC behavior while allowing their precise monitoring and manipulation. Here, we establish an in vitro culture system based on a 3D extracellular matrix environment and defined soluble factors, which for the first time allows expansion and long-term maintenance of murine multipotent HFSCs in the absence of heterologous cell types. Strikingly, this scheme promotes de novo generation of HFSCs from non-HFSCs and vice versa in a dynamic self-organizing process. This bidirectional interconversion of HFSCs and their progeny drives the system into a population equilibrium state. Our study uncovers regulatory dynamics by which phenotypic plasticity of cells drives population-level homeostasis within a niche, and provides a discovery tool for studies on adult stem cell fate. Synopsis An advanced in vitro culture system allows for enrichment and long-term maintenance of multipotent mouse hair follicle stem cells (HFSCs), recapitulating key features of their in vivo regulation. Combination of a 3D extracellular matrix environment and defined soluble components (FGF-2, VEGF-A, ROCK inhibitor Y27632) facilitates long-term propagation of HFSCs. HFSC expansion can be achieved from purified HFSCs as well as from total or HFSC-depleted epidermal cell mixtures. Cultured HFSCs retain multipotency, self-renewal potential, and transcriptional identity. Bidirectional interconversion of cultured HFSCs driven by BMP and Shh pathways leads to self-organization into a dynamic equilibrium between HFSCs and their progeny. Introduction Adult somatic stem cells (SCs) fuel tissue renewal, repair, and remodeling. The ability of SCs to tune their proliferation and differentiation rates to the changing needs of their resident tissues is central for the maintenance of organ homeostasis. Given their potency, even incremental alterations in SC behavior should lead to substantial changes in tissue size and architecture. Yet, such changes are strikingly rare, strongly implying that SCs are under tight homeostatic regulation allowing rapid adaptation of the system to disturbances to efficiently restore tissue functions. However, the mechanisms of such population-level regulation are poorly understood. Hair follicle stem cells (HFSCs) fuel cyclical bouts of adult hair follicle regeneration. Due to their well-defined quiescence-activation cycle, HFSCs represent an excellent paradigm for studying somatic adult SC lineage commitment (Blanpain & Fuchs, 2014). HFSCs are activated in a two-step process: First, quiescent HFSCs are activated to generate primed HFSCs that in a second step establish a pool of transit-amplifying cells (TACs; Greco et al, 2009; Hsu et al, 2014b). TACs are a transition state between SCs and their differentiated progeny, and their generation is a rate-limiting step in SC differentiation (Hsu et al, 2014b). SCs reside in spatially distinct microenvironments termed niches that consist of neighboring cells, extracellular matrix and signals derived from these compartments. Niches integrate signals to adjust SC behavior to the needs of organisms, to prevent SC depletion and at the same time restrict excessive SC expansion into the surrounding tissue (Morrison & Spradling, 2008; Blanpain & Fuchs, 2014; Scadden, 2014). Even though the critical importance of niches in SC regulation has been established, their complexity in mammals has prevented identification of the precise nature and combination of niche-derived signals, and hindered mechanistic studies of adult SC regulation. Interestingly, lineage tracing and ablation studies have demonstrated that HFSCs are dispensable for regeneration and that activated progeny re-populate the ablated SC niche to sustain hair regeneration (Hsu et al, 2011; Rompolas et al, 2013). This suggests that the niche instructs reprogramming of committed progenitors to a SC state, providing a mechanism to ensure robustness of tissue homeostasis. However, the mechanisms of this niche-directed reprogramming are completely unknown. Thus, there is a fundamental need to unravel the complex signaling circuitry governing HFSC identity and behavior and to define how the niche instructs HFSC homeostasis. One major obstacle to uncovering these fundamentals has been the lack of a system for ex vivo maintenance of HFSCs in the absence of other heterologous cell types and that also allows precise manipulation and monitoring of HFSC fate decisions. While various 2D cell culture systems for epidermal keratinocytes exist (Barrandon & Green, 1987; Trempus et al, 2003; Blanpain et al, 2004; Jensen et al, 2010; Bilousova & Roop, 2013), methods to maintain and expand bona fide multipotent HFSCs in culture in the absence of feeder cells are lacking, as are in vitro methods to capture the dynamic behavior of HFSCs and their progeny. In the current study, we identify a specific combination of niche factors that for the first time allow expansion and long-term maintenance of HFSCs. Utilizing this system, we uncover self-organizing phenotypic plasticity and dynamic bidirectional interconversion between HFSCs and their progeny, providing a cellular mechanism for homeostatic regulation of a SC niche. Results Establishment of a HFSC culture system We aimed at reconstituting the essential components of the HFSC niche in vitro by applying knowledge gained from in vivo studies on signaling within the HFSC niche. Freshly isolated epidermal cells from telogen-stage mice (P21) contained 5.6 ± 1.2% (± SD) CD34+α6+ HFSCs (Fig 1A). These isolated epidermal cell suspensions were subsequently cultured in standard 2D culture conditions either in a keratinocyte growth medium (KGM) or in FAD medium on a fibroblast feeder layer, which are widely used culture conditions for murine keratinocytes (Watt & Green, 1982; Morgner et al, 2015). Flow cytometry analyses of cells grown under these conditions demonstrated that the CD34+α6+ HFSC population was depleted within 14 days (Fig 1A and B, and Appendix Fig S1A). Figure 1. Establishment of a HFSC culture system Representative FACS plots of freshly isolated epidermal cells (Epi d0) and day 14 (d14) cultures of epidermal cells in different growth conditions and media. KGM 2D: 2D in basal KGM; 3C: 3D-Matrigel in KGM + Y27632 + VEGF-A + FGF-2. CD34+α6+ cells were quantified by flow cytometry from d14 as in panel (A) (mean ± SD; n = 3–5; *P ≤ 0.05; ***P ≤ 0.001, Kruskal–Wallis/Dunn's post-test). 3C 2D: 2D in KGM + Y27632 + VEGF-A + FGF-2; Y: 3D-Matrigel in KGM + Y27632; YV: Y + VEGF-A; YF: Y + FGF-2; 3C: Y + VEGF-A + FGF-2. Absolute numbers of CD34+α6+ cells from Epi d0 and d14 cultures as in (A and B). Fold enrichment over Epi d0 is shown (mean ± SEM, n = 3–5; *P ≤ 0.05, Mann–Whitney U-test). Epidermal cells grown for 14 days in the indicated conditions. 3C: 3D-Matrigel in KGM + Y27632 + VEGF-A + FGF-2. 3C 2D: 2D in 3C medium. Scale bars 30 μm. Immunofluorescence analysis of 3C cultures showing the expression of the HFSC markers CD34 and SOX9 in a subset of cells. Scale bars 25 μm. See also Appendix Fig S1. Download figure Download PowerPoint As laminins are important for HFSC maintenance (DeRouen et al, 2010; Morgner et al, 2015), and laminin-rich basement membrane extracts such as Matrigel have been successfully used to support growth of other epithelial SCs (Sato & Clevers, 2013), we tested whether providing a basement membrane-like extracellular matrix 3D environment would support HFSC growth in culture. We embedded freshly isolated epidermal cells in Matrigel and cultured them in KGM. As cells failed to survive under these conditions, we included a ROCK inhibitor (Y27632) to prevent anoikis that occurs due to deprivation of cell–cell contacts (Hofmann et al, 2007). These culture conditions provided a small but consistent increase in the population of CD34+α6+ HFSCs compared to cells cultured in 2D (Fig 1B). Several growth factors, including FGFs, EGF and VEGF-A, are expressed in cells of the HF and have been shown to regulate their growth (Kozlowska et al, 1998; Ozeki & Tabata, 2002; Doma et al, 2013; Plikus & Chuong, 2014). Combining Y27632 with either FGF-2 or VEGF-A in KGM-3D Matrigel resulted in a further relative increase in CD34+α6+ cells (Fig 1B), but these media did not increase the absolute numbers of CD34+α6+ cells (Fig 1C), suggesting that both conditions promoted HFSC survival and/or growth to a limited extent. In contrast, culturing epidermal cells in KGM-3D Matrigel containing Y27632, FGF-2, and VEGF-A (from now on referred to as 3C) significantly expanded CD34+α6+ cells within the cultured keratinocyte population (Fig 1A–C), inducing a ~sevenfold increase in the relative amount and a fivefold increase in absolute numbers of CD34+α6+ cells after 14 days of culture (Fig 1A–C). FGFs 7, 10, and 18 promoted enrichment of CD34+α6+ cells to the same extent as FGF-2 (Appendix Fig S1B). The 3D configuration was essential, as there was no enrichment of CD34+α6+ cells when cells were cultured in the 3C medium in standard 2D conditions (3C 2D; Fig 1B). Interestingly, the commonly used keratinocyte medium FAD supplemented with 3C failed to support cell growth even in the presence of 3D Matrigel (Appendix Fig S1C). Next, we analyzed the morphology of cells grown in 3C conditions. Cultures containing CD34+α6+ cells grew into spheroid cell clusters that did not generate a lumen (Fig 1D), in contrast to other epithelial-cell 3D culture systems (Lee et al, 1984; Sato et al, 2009). Interestingly, the complex laminin-rich mixture of matrix proteins present in Matrigel was dispensable as CD34+α6+ cells could also be grown in type I collagen gels using 3C medium (Appendix Fig S1D), most likely due to the expression and deposition of a laminin-332 meshwork that was observed in both Matrigel and collagen I cultures (Appendix Fig S1E). Immunofluorescence analysis of 3C cultures revealed expression of the HFSC markers CD34, keratin-15, and SOX9 (Fig 1E and Appendix Fig S1F and G), the latter a pioneer transcription factor required to establish HFSC identity (Adam et al, 2015). Expression of these markers was not detected in 2D cultures even in the presence of 3C medium (Appendix Fig S1G). Importantly, the cultures comprised of 97.7 ± 0.63% epithelial cells as indicated by staining for the epithelial cell adhesion molecule (EpCAM; Appendix Fig S1H). Altogether, these data show that a 3D environment together with FGF-2, VEGF-A, and Y27632 provide an essential set of HFSC-niche factors that are both necessary and sufficient to enable growth and expansion of cells with HFSC characteristics. Cells in 3C cultures retain self-renewing capacity and multipotency To assess whether cells cultured in 3C conditions maintain their proliferative potential characteristic for HFSCs, colony-forming assays were performed (Jensen et al, 2010). Cells from 3C cultures subsequently plated on feeders at clonal density gave rise to more colonies that were also larger in size compared to freshly isolated epidermal cells containing 5.6 ± 1.2% HFSCs (Fig 2A). In addition, these colonies contained small, tightly packed, cobblestone-like colonies (Appendix Fig S2A), characteristic of holoclones observed in feeder-dependent 2D cultures of purified HFSCs (Blanpain et al, 2004; Greco et al, 2009). These findings indicate that the 3C conditions enrich for epidermal cells with high proliferative potential, a hallmark of HFSCs and their immediate progeny (Blanpain et al, 2004; Greco et al, 2009). Figure 2. Cells in 3C cultures retain self-renewing capacity and multipotency Colony-forming assays show increased proliferative potential of cells cultured in 3C compared to freshly isolated epidermal cells (Epi d0; control; mean ± SEM; n = 5; *P ≤ 0.05, Mann–Whitney U-test). 3C: 3D-Matrigel in KGM + Y27632 + VEGF-A + FGF-2. Full-thickness skin reconstitution assay with freshly isolated epidermal cells (Epi d0) or 3C cultures shows that cells cultured in 3C retain their self-renewal capacity and multipotency. A representative recipient of four mice/condition is shown. Right panel shows quantification of hair follicle density. Note that 3C cultures give rise to more hair compared to controls (mean ± SEM; n = 4; *P ≤ 0.05, Mann–Whitney U-test). Long-term culture of cells in 3C was performed as shown in the left panel. 3C cultures maintain a stable population of CD34+α6+ long term (data are shown using a box-and-whisker plot: box indicates 25th and 75th percentiles; error bars represent 10th and 90th percentiles; middle line is the median; n = 3–5 independent experiments). p: passage. Full-thickness skin reconstitution assay using 3C cells from passage 1 (p1) and passage 15 (p15) shows that cells cultured in 3C maintain their multipotency and self-renewal capacity in long-term cultures. A representative recipient of four mice/condition is shown. See also Appendix Fig S2. Download figure Download PowerPoint In order to evaluate whether the ex vivo expanded CD34+α6+ cells represented functional HFSCs, we examined their self-renewal and multipotency in vivo. To this end, we performed full-thickness skin reconstitution assays (Lichti et al, 1995; Blanpain et al, 2004; Jensen et al, 2010). Cells cultured in 3C fully reconstituted the epidermis and produced hair, and as expected, were slightly more efficient in their regenerative capacity compared to freshly isolated epidermal cells reflecting the difference in HFSC quantity (5.6 ± 1.2% CD34+α6+ in epidermis compared to 42 ± 7% CD34+α6+ in 3C cultures; Fig 2B). This demonstrates that cells cultured in 3C preserve their multipotency and capacity to self-renew. Strikingly, 3C cultures could be passaged every 2 weeks into fresh 3D Matrigel for a period of at least 20 weeks without changes in the levels of CD34+α6+ cells (Fig 2C) or their proliferative potential (Appendix Fig S2B). The proportion of CD34+α6+ cells within the 3C cultures remained largely stable from the first passage onward and the cells propagated long-term were still fully competent to generate hair in full-thickness skin reconstitution assays (Fig 2D). Moreover, freeze–thaw experiments demonstrated that 3C cultures could be stored frozen and cultured again without loss of the CD34+α6+ population or its ability to generate hair (Appendix Fig S2C and D). Collectively, these data demonstrate that the 3C conditions recapitulate the conditions of a SC niche, enabling maintenance and enrichment of functionally competent cells that display HFSC characteristics. In addition, long-term 3C cultures sustain a stable equilibrium between CD34+α6+ and CD34−α6+ cells that can be stored frozen without loss of multipotency. Transcriptomes of cells cultured in 3C resemble in vivo HFSCs In order to provide an optimal model to uncover novel HFSC biology, cells cultured in 3C should share high resemblance to in vivo HFSCs also on the molecular level. To examine the molecular identity of cells cultured in 3C, we performed RNA sequencing (RNAseq) and compared the transcriptomes of epidermal cells cultured in 3C to freshly isolated epidermal cells that were used to establish the 3C cultures as well as to FACS-purified in vivo CD34+α6+ HFSCs (Fig 3A and B; Dataset EV1). 3C cultures more closely resembled in vivo HFSCs than the epidermal cell mixtures they were derived from, as shown by Euclidian distance (Fig 3B), Pearson's correlation, and principal component analysis (Appendix Fig S3A and B). Figure 3. Transcriptomes of cells cultured in 3C resemble in vivo HFSCs Schematic workflow of the RNAseq experiment from FACS-purified CD34+α6+ HFSCs (CD34+α6+), cells cultured in 3C and freshly isolated epidermal cells (Epi d0). Heatmap and Euclidian distance dendrogram of quantified transcripts from RNAseq data generated as shown in panel (A). 3C cultures cluster with purified CD34+α6+ HFSCs (n = 3 biological replicates). Venn diagram of genes most significantly upregulated in 3C and in purified CD34+α6+ (log2FC > 2, padj < 0.05) together with published HFSC signature shows significant overlap between the three signatures (P = 0.00252451 and P = 3.058928e-51, hypergeometric distribution). Schematic workflow of the RNAseq experiment from FACS-purified CD34+α6+ and CD34−α6+ cells from 3C cultures. Heatmap of selected genes differentially expressed in 3C-CD34+α6+ and 3C-CD34−α6+ cells. Note upregulation of multiple key HFSC lineage identity transcription factors and bulge markers (triangles) in 3C-CD34+α6+ cells, and upregulation of genes implicated in lineage progression in 3C-CD34−α6+ cells (circles). Venn diagram of genes upregulated in 3C-CD34+α6+ and in purified CD34+α6+ cells (log2FC > 2, padj < 0.05) together with a published HFSC signature shows significant overlap between the three groups (P = 1.381896e-09 and P = 0.01636903, hypergeometric distribution). RT–qPCR analyses of FACS-purified CD34+α6+ and CD34−α6+ cells from either 3C cultures or from freshly isolated epidermis show upregulation of HFSC identity genes in both freshly isolated and 3C-CD34+α6+ cells compared to freshly isolated CD34−α6+ progenitors. 3C-CD34−α6+ cells show intermediated expression (mean ± SEM; n = 4; *P ≤ 0.05, **P ≤ 0.01, Kruskal–Wallis). Both 3C-CD34+α6+ and 3C-CD34−α6+ cells show upregulation of a panel of cell cycle genes and genes shown to be enriched in HFSC progeny (mean ± SEM; n = 4; *P ≤ 0.05, **P ≤ 0.01, Kruskal–Wallis). See also Appendix Fig S3 and Datasets EV1, EV2, and EV3. Download figure Download PowerPoint We next examined in which respects the gene expression profiles of 3C cultures resembled and differed from in vivo HFSCs. For this, we computed genewise Euclidian distance calculations of the three conditions (Epi d0, 3C, CD34+α6+ HFSCs) to identify clusters of genes that explained most of the variance in gene expression among the three groups. Gene ontology (GO) term analysis of the top 5 clusters revealed three clusters of genes (clusters 1, 2, and 5; Appendix Fig S3C) that showed comparable, lower expression levels in 3C and CD34+α6+ HFSCs and higher expression levels in Epi d0 (Appendix Fig S3C). These clusters contained GO terms for protein translation, protein transport, and metabolism. This was intriguing, as low protein translation rates have been recently linked to HFSC identity and function (Blanco et al, 2016). Cluster 3 contained genes that displayed an intermediate expression level in 3C (Appendix Fig S3C). These genes were enriched in GO terms for metabolism, translation, and oxidative phosphorylation. Only one cluster (cluster 4; Appendix Fig S3C) was identified for genes that were different in 3C when compared to CD34+α6+ HFSCs, which enriched for genes involved in transcription, tRNA processing, and DNA replication, possibly reflecting the slower replication rates of the quiescent CD34+α6+ HFSCs isolated from telogen-stage mice compared to cells in culture. We further analyzed the genes that were similarly expressed in 3C cultures and purified CD34+α6+ HFSCs in comparison with total epidermal cells in more detail. The most significantly upregulated genes (log2FC > 2, padj < 0.05; Dataset EV2) were compiled to generate gene expression enrichment profiles for 3C cultures and purified CD34+α6+ HFSCs, respectively, which were then compared to each other and to a compiled HFSC signature from published datasets (Morris et al, 2004; Tumbar et al, 2004; Lien et al, 2011; Dataset EV2). Despite being a ~50:50 mixture of CD34+α6+ and CD34−α6+ cells, genes enriched in 3C cultures showed significant overlap with genes enriched in purified CD34+α6+ HFSCs (P = 3.058928e-51) and, importantly, with the published HFSC signature (P = 0.00252451) including key HFSC lineage identity genes such as Cd34, Sox9, Tcf3 (Fig 3C). This provided further support that cells propagated in 3C cultures resemble HFSCs. We proceeded to explore the differences in gene expression between 3C cultures and purified CD34+α6+ HFSCs. GO term and Gene set enrichment (GSEA; (Subramanian et al, 2005) analyses revealed that genes involved in Wnt signaling, regulation of cell proliferation, extracellular matrix organization, and keratinocyte differentiation were among the most significantly overrepresented among the differentially expressed genes (Appendix Fig S3D–F). Closer examination of the specific genes revealed that negative regulators of Wnt such as Dkk3 and Sfrp1, as well as Wnt ligands shown to be regulated by the BMP signaling pathway (Wnt 5b, 16), were up in CD34+α6+ HFSCs whereas Wnts shown to be enriched in activated HFSCs or their progeny (Wnt 4, 9a, 10a, 10b) were upregulated in 3C (Appendix Fig S3F). Furthermore, positive regulators of the cell cycle such as cyclin D1 and G1 (Ccnd1 and Ccng1), as well as genes associated with keratinocyte differentiation were upregulated in 3C (Appendix Fig S3F). A number of ECM genes were upregulated in CD34+α6+ HFSCs possibly reflecting the excess of ECM proteins present in the Matrigel of the 3C cultures (Appendix Fig S3E and F). As the transcriptome-wide expression analyses indicated that cells cultured in 3C conditions closely resembled in vivo HFSCs, but that differences in Wnt and BMP signaling pathway regulators and targets as well as cell cycle regulators were observed, we hypothesized that the 3C cultures consist of a ~50:50 mixture of HFSCs (CD34+α6+ cells) and their progeny (CD34−α6+ cells). To test this, we FACS-purified CD34+α6+ and CD34−α6+ cells from 3C cultures and performed RNAseq (Fig 3D and E; Dataset EV3). Remarkably, 47% of the genes upregulated in CD34+α6+ cells from 3C cultures (3C-CD34+α6+) were components of HFCS signatures (Fig 3E and F), including several key lineage identity genes such as Lhx2, Id2, Sox9, Nfatc1, Tcf3, as well as bulge markers Krt15 and Cd34 (Fig 3E). In contrast, genes downregulated in the 3C-CD34+α6+ population contained markers of early lineage commitment (Fig 3E and Appendix Fig S3I), previously shown to be upregulated in HFSC progeny (TACs, hair germ or hair matrix cells; Greco et al, 2009; Lien et al, 2011). To validate the sequencing data and to dissect the gene expression differences in 3C cells (3C-CD34+α6+ and 3C-CD34−α6+) and freshly isolated in vivo CD34−α6+ (basal progenitors) and CD34+α6+ HFSCs in more detail, we performed RT–qPCR analyses. Importantly, these analyses corroborated the RNAseq results and demonstrated that key HFSC lineage identity genes (Sox9, Tcf3, Lhx2, Nfatc1, Id2, and Dkk3) were highly enriched in 3C-CD34+α6+ compared to progenitors in vivo (CD34−α6+) or in 3C (3C-CD34−α6+), and in most cases expressed at comparable levels as in in vivo CD34+α6+ HFSCs, confirming that the 3C-CD34+α6+ cells are HFSCs (Fig 3G). In line with the RNAseq data, a panel of cell cycle regulators and hair germ/matrix markers showed upregulation in both 3C populations compared to freshly isolated cells (Fig 3H), most likely reflecting their active expansion compared to the telogen-stage in vivo CD34+α6+ cells (Fig 1C). Interestingly, 3C-CD34−α6+ cells showed mild upregulation of early lineage commitment genes (Fig 3E and Appendix Fig S3G), but displayed higher expression of the key HFSCs lineage identity genes compared to in vivo CD34−α6+ cells (Fig 3G), suggesting that these cells could represent immediate progeny of 3C-CD34+α6+ cells. 3C cultures evolve into a stable population equilibrium of HFSCs and non-HFSCs Our data so far showed that long-term 3C cultures established and maintained a stable mixture of CD34+α6+ HFSCs and CD34−α6+ progenitors (Fig 2C). To understand how this was generated, we first analyzed the dynamic evolution of the population equilibrium in 3C cultures generated from freshly isolated epidermal cell mixtures. Time course analyses revealed that after an initial drop in both CD34+α6+ and CD34−α6+ cell numbers, both populations steadily expanded starting from day 2 with CD34−α6+ showing a more rapid increase (Fig 4A). However, after both populations reached peak values by day 8, the numbers of CD34−α6+ declined to establish an approximate 50:50 equilibrium of HFSCs and non-HFSCs (Fig 4A). This was accompanied by a plateau in the size of the 3C cell clusters (Appendix Fig S4A). The population equilibrium remained constant during passaging (Appendix Fig S4B), similar to what was observed in long-term cultures (Fig 2C). Interestingly, determination of proliferation rates by EdU incorporation after the cultures had reached equilibrium (d10-d14) showed that cultured CD34+α6+ cells cycled slightly faster than the CD34−α6+ population (Fig 4B), whereas the apoptosis rates were comparable between the two populations (Appendix Fig S4C). Figure 4. 3C cultures evolve into a stable population equilibrium of HFSCs and non-HFSCs FACS analysis of CD34+α6+ cells over time shows that 3C cultures reach a ˜50:50 population equilibrium after 10 days of culture (mean ± SEM; n = 3). Cells were EdU-labeled for 24 h (day 10–11) prior to analysis by flow cytometry to quantify proliferating cells. Percentage of EdU+ cells is indicated (n = 4; mean ± SD; *P ≤ 0.05, Mann–Whitney U-test). Schematic illustration of the parameters used in the mathematical model. HFSCs (CD34+α6+) and non-HFSCs (CD34−α6+) have a maximum proliferation rate of pSC,max and pP,max, respectively. They enter apoptosis with rates apSC and apP and are both degraded with a rate deg. Non-HFSCs can convert to HFSCs with a rate tP→SC and vice versa with rate tSC→P. Simulated population proportions of CD34+α6+ HFSCs and CD34−α6+ non-HFSCs d
Referência(s)