Hair Follicle Development in Mouse Pluripotent Stem Cell-Derived Skin Organoids

Jiyoon Lee et al.
2018-01-02

Highlights
•Skin organoids can be generated from mPSCs under defined conditions
•Skin organoids are composed of self-assembled epidermal and dermal layers
•Skin organoids produce hair follicles, sebaceous glands, and adipocytes
•Hair follicle induction from skin organoids mimics normal hair folliculogenesis
Summary
The mammalian hair follicle arises during embryonic development from coordinated interactions between the epidermis and dermis. It is currently unclear how to recapitulate hair follicle induction in pluripotent stem cell cultures for use in basic research studies or in vitro drug testing. To date, generation of hair follicles in vitro has only been possible using primary cells isolated from embryonic skin, cultured alone or in a co-culture with stem cell-derived cells, combined with in vivo transplantation. Here, we describe the derivation of skin organoids, constituting epidermal and dermal layers, from a homogeneous population of mouse pluripotent stem cells in a 3D culture. We show that skin organoids spontaneously produce de novo hair follicles in a process that mimics normal embryonic hair folliculogenesis. This in vitro model of skin development will be useful for studying mechanisms of hair follicle induction, evaluating hair growth or inhibitory drugs, and modeling skin diseases.

Introduction
The integumentary system consists of skin and its appendages. The skin is composed of two layers, the epidermis and the dermis, which produce appendages, such as hair follicles (HFs), sweat glands, and nails. While the epidermis of the skin arises from the ectoderm, the dermis has different embryonic origins. The majority of the body dermal tissue arises from paraxial and lateral plate mesoderm, while facial dermis arises from cranial neural crest cells (Dequéant and Pourquié, 2008, Driskell and Watt, 2015, Fernandes et al., 2004). Regardless of dermal origin, all skin types require the interplay of epithelial (epidermis) and mesenchymal (dermis) cells for terminal development and appendage formation. In animal models, rodent and human skin with HFs can be reconstituted by co-culturing mesenchymal cells with epithelial cells or, specifically, HF-initiating dermal papilla (DP) cells (Asakawa et al., 2012, Chuong et al., 2007, Ehama et al., 2007, Ikeda et al., 2009, Nakao et al., 2007, Takagi et al., 2016, Toyoshima et al., 2012, Zheng et al., 2005). However, a chemically defined means of generating folliculogenic skin from pluripotent stem cells (PSCs) in vitro has been intangible.

To date, in vitro skin derivation strategies have focused on generating keratinocytes and fibroblasts from PSCs in separate cultures first and then combining the two types of cells to form a skin-like bilayer (Gledhill et al., 2015, Itoh et al., 2013, Oh et al., 2013, Sun et al., 2014). Recently, we developed a three-dimensional (3D) mouse embryonic stem cell (mESC) culture capable of producing cranial surface epithelia (also known as non-neural ectoderm), a precursor tissue of the skin epidermis. In the process of culture, a heterogeneous population of mesenchymal and neural cells is also generated (Koehler and Hashino, 2014, Koehler et al., 2013). Using this technique, we have demonstrated how to generate inner ear organoids that contain sensory epithelia reminiscent of postnatal mouse vestibular organs (Koehler et al., 2013, Koehler and Hashino, 2014). In addition, we also briefly described how the induced surface epithelia generate p63+ keratin 5 (KRT5)+ basal keratinocyte-like cells that self-organize into a cyst (Koehler et al., 2013), referred to hereafter as a skin organoid. In this article, we report that the mouse PSC-derived skin organoids recapitulate key steps of integumentary development and have the capacity to generate de novo HFs. Our results support an approach to generate skin and skin appendages from PSCs wherein the epidermal and the dermal cells are co-induced in a single organoid unit.

Results
Modulating TGF, FGF, and BMP Signaling Pathways Initiates HF Formation in R1 mESC Aggregates
In our previous studies, we described a cell culture treatment regimen capable of inducing surface ectoderm in aggregates of mouse PSCs (Koehler et al., 2013, Koehler and Hashino, 2014). An initial treatment with a transforming growth factor β (TGF-β) inhibitor, SB431542 (SB), and recombinant BMP4 (BMP) specifies a surface ectoderm at the outermost region of the spheroid cell aggregates. A subsequent treatment with FGF-2 (FGF) and a BMP inhibitor, LDN-193189 (LDN), promotes induction of placodal epithelium. PAX8 is expressed in patches of the epithelium by day 8 (Koehler et al., 2013), suggesting induction of cells similar to those in a cranial region that produces epidermis as well as otic and epibranchial placode derivatives (e.g., the inner ear and sensory neurons in cranial nerves VII, VIII, and IX; Groves and Fekete, 2012). While the epithelium develops on the aggregate surface, PSCs persist in the aggregate core during the first week of differentiation (DeJonge et al., 2016). In addition, an intermediate tissue layer between the PSC core and the surface ectoderm forms with Brachyury+ mesendoderm cells and N-cadherin+ SOX2+ neuroectoderm cells (Koehler et al., 2013). During inner ear organoid production, otic vesicles evaginate from the epithelium, and the intermediate layer comprised of mesoderm/neuroectoderm cells erupts outward to form an outer layer, leaving the surface ectoderm inside as an epidermal mantle (inside-out morphology; Figure 1A). While investigating the otic culture system using the R1 mESC line, we observed the occasional formation of protruding bulb-like structures that differed in morphology from inner ear organoids (Figure 1B). We discovered that these structures were extensions of the KRT5+ epidermis and were associated with SOX2+ DP-like cells (Figures 1C and 1D), reminiscent of nascent guard, awl, or auchene HFs (Driskell et al., 2009, Lesko et al., 2013). This observation was made independently by the Koehler and Heller laboratory groups. In light of these unexpected findings, we sought to better characterize the process of folliculogenesis.

HF Formation Is Cell-Line and Aggregate-Morphology Dependent
The R1 cell line displayed relatively low frequency of HF production (18% ± 8% of aggregates with HF-like bulbs per batch, n = 84 organoids, 6 experiments; Table S1A). Surprisingly, while testing our otic induction method on other cell lines, we noted that an Atoh1/nGFP mESC line (Lumpkin et al., 2003, Oshima et al., 2010) was highly folliculogenic (Figure 2; Table S1A). As reported previously, this Atoh1/nGFP mESC line rarely produced inner ear organoids without an additional treatment with a GSK3β inhibitor, CHIR99021 (CHIR), on day 8 of differentiation (DeJonge et al., 2016). Without CHIR treatment, however, we found that Atoh1/nGFP cells produced a significantly higher percentage of HF-bearing aggregates than R1 aggregates (Atoh1/nGFP: 83% ± 5% of aggregates with HF-like bulbs per batch, n = 193 organoids, 10 experiments, p < 0.0001; Table S1A). To uncover differences between the cell lines, we examined the events leading to induction of skin organoids. Similar to the R1 cell line, we observed induced Brachyury+ mesendodermal cells and ECAD+ PAX8+ epithelial cells during days 5–8 of differentiation of Atoh1/nGFP cells (Figure S1). By day 14, KRT5 was expressed in the epithelium in both R1 and Atoh1/nGFP organoids (n = 6 aggregates per cell line, 3 experiments; Figures 2A and 2B). Each aggregate typically contained a single skin organoid unit with KRT5 expressing cells forming a continuous spherical mantle-like structure (Figures 2A and 2B). While R1 cell aggregates appeared to have more pronounced growth of non-epithelial tissues around the KRT5+ epithelium, such as neuroectoderm cells denoted by SOX2 expression (Figures 2A and S2A), most Atoh1/nGFP aggregates either lacked or had one patch of non-epithelial tissue attached to the skin organoid during days 12–18 (Figures 2B and S2A). Based on this observation, we classified day 18 Atoh1/nGFP aggregates into three categories to reflect the degree to which the epithelium was covered by exterior non-epithelial tissues: uncovered, partially covered, and fully covered (see Figure 2C, representative images). Moreover, we grouped HF-bearing organoids into three categories based on the number of HFs they produced: 1–5, 6–15, or >15 HFs (see Figure 2D, representative images). Aggregates in each category, which were sorted by the amount of exterior non-epithelial tissue, produced HFs (Figure 2E, center pie graph). However, uncovered or partially covered aggregates produced the highest percentage of skin organoids bearing >15 HFs, while fully covered Atoh1/nGFP aggregates, similar to R1 aggregates, rarely produced organoids with more than five HFs (Figure 2E, branched pie graph). Based on these observations, we questioned whether HF production is a unique predisposition of the R1 and Atoh1/nGFP cell lines. To answer this question, we cultured a C57BL/BJ induced PSC (iPSC) line using the same skin organoid induction protocol. The C57BL/BJ iPSCs also generated HF-bearing skin organoids (Figure S2B), suggesting that robust induction of HF-producing skin organoids is not limited to one cell line. Moreover, as Atoh1/nGFP aggregates, uncovered or partially covered C57BL/BJ iPSC aggregates produced higher number of HFs compared to fully covered aggregates bearing only 0–2 HFs (data not shown). Together, these findings suggest that PSC line-specific characteristics affect the efficacy of HF production, whereas the general ability to produce HFs was observed in all three PSC lines.

Optimal Treatment Regimen for HF Induction
We next asked whether the complete SB, BMP, FGF, and LDN treatment regimen was necessary for HF induction. We knew from our previous work that SB/BMP-treated aggregates, without subsequent FGF and LDN treatment, produced epidermal keratinocyte cysts largely devoid of surrounding mesenchymal tissue (Koehler et al., 2013). Upon reexamination, we confirmed that HFs never developed when aggregates were grown in SB/BMP conditions (n = 12 aggregates, 3 experiments). Likewise, SB/BMP-FGF-treated aggregates produced epidermal cysts but were not folliculogenic (n = 12 aggregates, 3 experiments). Interestingly, SB/BMP-LDN-treated aggregates generated HF-bearing skin organoids, albeit inconsistently from batch to batch (46% ± 13% of organoids with HF-like bulbs per batch, n = 61 organoids, 8 experiments). However, we noted that the percentage of aggregates generating 15 or more HFs was significantly greater under the SB/BMP-FGF/LDN full-treatment condition compared to SB/BMP-LDN conditions (Table S1B; 45% ± 7% [SB/BMP-FGF/LDN] versus 13% ± 7% [SB/BMP-LDN] of organoids, respectively; p < 0.01). Thus, the full treatment of SB/BMP-FGF/LDN appears to be optimal for HF-bearing skin organoid formation. Moreover, when we treated the aggregates with CHIR on day 8, as used previously to encourage inner ear organoid induction (DeJonge et al., 2016), we found that presumptive HFs formed alongside inner ear organoids (Figure S2C). Consequently, the pulse of GSK3β inhibition does not disrupt HF induction. Self-Assembly of Skin Organoids Recapitulates Embryonic Development Using embryonic stem cell (ESC)- and iPSC-derived aggregates, we examined the integumentary developmental stages represented in our culture system (see schema in Figure 3A). In the developing murine skin, basal keratinocytes that express KRT5 are first observed in the surface epithelium at approximately embryonic day 9.5 (E9.5). Subsequently, intermediate or spinous keratinocytes that express KRT10 arise at ∼E14.5. Finally, the granular and cornified epidermal layers that express loricrin and filaggrin (FLG) arise at ∼E18.5 (Figure 3A). Remarkably, skin organoids develop stratified epidermis (Figures S3A and S3A′). We first observed KRT5 expression, indicative of basal keratinocytes, in patches of the ECAD+ TFAP2α+ epithelium on day 10 of differentiation (Figure 3B and S3B). We then noticed KRT10 expression in spinous-like epidermal cells by day 14 of differentiation (Figure 3C). Later, at day ∼25, we observed robust expression of loricrin and FLG in granular and cornified layers of the organoid epidermis, respectively (Figures 3D-3E′ and S3B′). Interestingly, we noted that these mature epidermal markers were typically absent from sections of the epithelium abutting non-epithelial cells, such as TUJ1+ neurons, suggesting that the composition of non-epithelial tissues may affect keratinocyte differentiation (Figures 3D–3E′ and S3B′). Concurrently with epidermal layer development, a dermal layer was induced on the surface of skin organoids. In the developing mouse skin, the dermis stratifies to form three distinct collagen-rich layers during development. At ∼E12.5, the dermis is composed of homogeneous dermal fibroblast progenitors, which later differentiates to form papillary and reticular dermal layers (E18.5), followed by an adipocyte-rich hypodermal layer (P2; Driskell et al., 2013). In our culture system, we noticed the presence of collagen type III, alpha-1 chain (COL3α1) and collagen type IV, alpha-1 chain (COL4α1) by day 12 of differentiation (Figure S3C); these are known extracellular matrix (ECM) proteins localized at the epidermal-dermal interface (Figures 3F′ and 3G′). The COL4α1 present on day 12 may be from Matrigel included in the medium on day 8; however, COL4α1 persisted throughout differentiation, suggesting that it may be produced endogenously (Figure 3G′ and S3C). Moreover, COL3α1, which is not a component of Matrigel, is also present at the epidermal-dermal interface beginning on day 12 and was present throughout differentiation (Figure 3F′ and S3C). By day 14 of differentiation, a dermal fibroblast-like cell marker, DLK1, was detected in cells around the epidermis (Figure 3F). As the skin organoids differentiate (from day 16), a DLK1+ fibrotic region emerged in the dermal layer. Likewise, CD34, another fibroblast-like cell marker, was detected from day 12 surrounding the epidermis (Figure 3G). Interestingly, by day 16, CD34+ cells were present throughout the dermal layer, while DLK1+ cells were localized to discrete regions, suggesting that a diverse population of dermal cells may emerge in skin organoids (Figures 3F and 3G), consistent with late embryonic and early postnatal dermal development (Driskell et al., 2013). CD34 was expressed in the dermal layer at later stages of differentiation as well (day 26; Figure S3C′). Pearl-shaped adipocytes started to appear on day 26 (Figures 3H–3I′), mimicking the maturation of fat in the hypodermis in vivo. Thus, the mouse skin organoids produced in our culture system recapitulate key features of skin differentiation on a timescale that roughly correlates with normal embryonic development. Skin Organoid HF Induction Mimics Early Embryonic Developmental Stages In mice, HF development has been classified into discrete stages (Paus et al., 1999; Figure 4A). During stage 1, at the site of HF induction, dermal papillary fibroblasts form a condensed mass as the epidermis thickens and protrudes to generate the hair germ. In stages 2–3, rapidly dividing Ki67+ epithelial cells elongate the germ into a peg-like structure with DP cells at the terminal end. In stages 4–6, the hair pegs continue to elongate and thicken to form a bulb, cell layers of the inner and outer root sheaths become defined by morphology and expression of specific protein markers, and the hair cortex and shaft forms. In our in vitro system, hair germs were visible as early as days 14–18 of differentiation by double staining with EDAR and LHX2 (Figures 4B, S4A, and S4A′). Subsequently, HFs reminiscent of early-stage embryonic HFs (stages 1–4; Figure 4A) could be identified in skin organoids at any time point between differentiation days 18–26 (Figures 4C–4D″), suggesting that new HF induction is continuous during this time period. By day 18 of differentiation, all of the developing HFs contained SOX2+ dermal condensates/DPs, indicating the development of nascent guard, awl, or auchene HFs (Figure 4D–4D″). Interestingly, however, at the later stage of differentiation by day 28, we also noticed the presence of HFs with SOX2-negative hair bulb region, reminiscent of zigzag HFs (Figures 4E-4E″, S4B, and S4B′). These findings suggest that skin organoids may be capable of producing at least two types of HFs—SOX2+ guard, awl, or auchene HFs and SOX2− zigzag HFs—in a single skin organoid unit. By dissecting the structural architecture of more mature-looking HFs after day 24 of differentiation, we discovered that the HFs generated within our skin organoids shared key features with HFs in embryonic stages 7–8. The HFs consisted of an outermost layer of αSMA+ dermal sheath (Figure 4F), KRT5+ p63+ outer root sheath (Figure 4G), GATA3+ inner root sheath layers (Figures 4H–4I′), Ki67+ p63+ HF matrix (Figure 4C, 4C′, and 4G), and AE13+ cortex cells in the HF shaft (Figures 4I, 4I′, and S4D″). The proper organization of the HF in the skin organoid was revealed by transmission electron microscopy (TEM), showing all of the HF lineages formed from the outermost dermal sheath to the center of the hair shaft medulla (Figures 4J–4J″). Together, these findings reveal that skin organoids produced in our culture system can generate HFs similar to those seen in late embryonic and early postnatal development, consisting of all major units, including HF matrix, inner root, outer root, and dermal sheath layers. Specialized Epidermal and Dermal Cellular Compartments Arise in Skin Organoids In addition to the basic HF components outlined above, our skin organoids are composed of other epidermal and dermal micro-niches. During embryonic stages 7–8, neural-crest-derived melanocytes in the HF matrix pigment the hair shaft, sebaceous glands (SGs), and arrector pili muscles develop, and the dermis forms a hypodermal layer rich with adipocytes (Paus et al., 1999). We investigated whether these key cellular niches of the pilosebaceous unit arise in skin organoids (Figure 5A). For these analyses, we utilized both floating and Matrigel-embedding methods for organoid cultures. In the floating culture, the HFs typically wrapped tightly around the organoid (Figure 5B; Movie S1), whereas the HFs grew outward in Matrigel droplets, making it easier to assess HF morphology (Figure 5B′). We noted that not all organoid HFs had normal morphology; each organoid we examined contained at least one abnormal HF, which will require further analysis to define abnormal characteristics (Figure 5B′, arrowheads). Restricting analysis to HFs with normal morphology, we identified ITGα8+ αSMA+ muscle-like structures with elongated nuclei in the organoid dermis (Figures 5C and 5C′), which could indicate arrector pili muscle induction (Driskell et al., 2013). However, we did not observe elongated ITGα8+ αSMA+ cells with attachment points on the HFs and interfollicular epidermis as would be expected for fully formed arrector pili muscles. After 24 days of differentiation, we could detect SG-like structures on organoid HFs by oil red O and LipidTOX staining (Figures 5D–5D″). The structures were located near the HF attachment point with the epidermal mantle and co-expressed ECAD and SCD1, consistent with SG identity (Figures 5E, 5E′, and S4C–S4C″). The majority of SG-like structures observed in the skin organoids were bifurcates, reminiscence of guard hairs (Driskell et al., 2009). Moreover, as mentioned earlier in Figure 4D′, the SOX2+ nuclei speckles in the skin epithelium were Atoh1/nGFP+ ISL1+ cells that arose in the epithelium between days 14 and 16 (Figures 5F–5F″), reminiscence of Merkel cells, which are touch-sensing cells in the epidermis (Lumpkin et al., 2003, Perdigoto et al., 2014). However, these cells were rarely seen after day ∼20 of differentiation (data not shown). We suspect that culture conditions or perhaps a lack of proper neuronal innervation may dysregulate Merkel cell development in the organoid culture format. Thus, further in-depth studies will be needed to additionally define our culture system for Merkel cell maturation. Remarkably, we also observed p75+ SOX10+ neural-crest-like cells in the mesenchymal layer (Figures 5G and S4D–S4E′; Movie S1). In addition, melanocytes were visible around the epidermis as early as day 21 (Figure 5H) and migrated to and pigmented HFs in a subset of Atoh1/nGFP mESC-derived skin organoids by day 26 (Figures 5H′, 5H″, and S4F), suggesting that melanocytes can develop from the neural-crest-like cell population. However, melanocyte induction was infrequent during production of Atoh1/nGFP skin organoid production (12/252 organoids; other cell lines were not quantified). Together, our data suggest that the development of key skin and HF accessory tissues is recapitulated in skin organoids, but some cellular compartments (e.g., melanocytes) may depend on the composition of non-epithelial tissue associated with skin organoids. Skin Organoid-Derived HFs Contain Bulge-like Regions It has been reported that slow-cycling HF stem cells reside in a bulge region of the outer root sheath located proximal to the SG (Cotsarelis et al., 1990, Lavker et al., 2003). To determine whether organoid HFs develop a bulge-like region with the follicular stem cells, we adapted an EdU (5-ethynyl-2′-deoxyuridine)-labeling assay. As EdU incorporates into newly synthesized DNA, it makes possible to distinguish proliferating cell populations. In our culture system, we observed robust EdU incorporation in HF matrix cells (Figure 6A; Movie S2), which corresponded with Ki67+ proliferating cells in the matrix (Figures 4C, C′, and 4G). EdU incorporation was less prevalent in regions of the HF above the matrix. Despite the sparse labeling, we consistently observed EdU incorporation in a subset of cells located in a bulge-like region near the SG in the upper outer root sheath (Figures 6A′ and 6A″; Movie S2). These cells may be hair follicle stem cell (HFSC)-like cells or SG progenitors. To further characterize cells in the bulge-like region, we examined known protein markers of HFSCs. We found that the HF bulge and epithelial stem cell markers, KRT15 and SOX9, were expressed broadly in the organoid HF outer root sheath as well as the bulge-like region (Morris et al., 2004, Nowak et al., 2008; Figures 6B–6B″). Nuclear-localized NFATc1 has been identified as a definitive marker of nascent and adult HFSCs, whereas CD34 is expressed in adult HFSCs of HFs that have entered the growth cycle (Horsley et al., 2008, Woo and Oro, 2011). We rarely observed KRT15+ follicular cells that co-expressed CD34, indicating that organoid HFs likely do not contain adult HFSC-like cells (Figures 6B-B″and S4G–S4H′). By contrast, NFATc1 was specifically expressed by cells in the presumptive bulge region of organoid HFs between days 26 and 30 (Figures 6C and 6C′). Notably, NFATc1 expression was predominantly localized to the cytosol in bulge region cells; however, we observed nuclear-localized NFATc1 expression in the bulge regions of some HFs (n = 2 HFs from 9 organoids, ∼40 HFs total; Figures 6C and C′). Based on previous studies, these data suggest that skin organoid HFs are capable of producing nascent HFSCs, reminiscent of HFSCs at a late embryonic (∼E17.5) stage of development (Horsley et al., 2008). The rarity of cells expressing nuclear NFATc1 may indicate that the current culture conditions are not optimal for maturation of the bulge region stem cell niche. Skin Organoid HFs Degenerate during Long-Term Culture Finally, we examined whether skin organoid HFs undergo a catagen-like degenerative process similar to HFs in vivo as they enter the growth cycle. We tracked individual HFs over time using differential interference contrast (DIC) imaging (Figure 6D). We found that the HFs continued to grow until the experiment was terminated on day 32 while the growth rates gradually decreased day by day, and eventually, the HF matrix deteriorated around day 32, implying the possibility that the HFs undergo catagen (Figures 6D and 6E). To analyze whether the HFs truly undergo HF cycling in our culture system, we extended the culture period to 50 days and monitored HF growth (3 additional experiments). We did not observe newly forming HFs beyond days 32–35. Between 35 and 50 days, HFs appeared to have arrested growth and a dense layer of adipocytes forms on the surface of the organoids. We noted that skin organoid HF shafts cannot shed as they do in vivo during a typical HF growth cycle, which may lead to abnormalities in skin organoid HFs transitioning from development stages into the growth cycle. Partial dissociation and re-plating organoids in Matrigel droplets did not improve skin organoid longevity (data not shown). Thus, further studies will be necessary to explore ways of increasing the longevity of skin organoid cultures to potentially support in vitro HF cycling. Discussion Our findings demonstrate that HF-bearing skin can be generated in vitro from a homogeneous source of mouse PSCs, under serum-free conditions. Not surprisingly, HF induction seems to be dependent on the co-development of both epidermal and dermal cells. In our culture system, the induction of epidermal cysts is highly reproducible across three PSC lines, yet the frequency of accompanying induction of dermis and HFs appears to be cell-line dependent. Cell-line variability is a commonly cited shortcoming of organoid systems that rely on tissue self-assembly (Chen et al., 2014, Völkner et al., 2016). For our skin organoid system, the heterogeneity of the exterior tissue mass that forms during days 8–12 of differentiation likely underlies variability in HF induction. In particular, our data suggest that non-dermal tissues may be physically and/or chemically inhibitory to HF induction in floating aggregates. For example, R1 aggregates, which rarely generated HFs, contained a high abundance of neural tissue. By contrast, folliculogenic Atoh1/nGFP mESC and C57BL/BJ mouse induced pluripotent stem cell (miPSC) aggregates contained less non-dermal tissue. Thus, restricting the composition and self-assembly of the dermal layer to exclusively cutaneous cell types (e.g., fibroblast, adipocytes, and melanocytes) will be a challenge for future studies on skin organoids. We found that minor modifications to the treatment concentrations and timing did not consistently improve or diminish the efficiency of HF induction. Thus, more targeted alterations must be made to gain control of dermis induction. One important consideration may be the use of physicochemically defined matrices, such as polyethylene glycol hydrogels, to replace Matrigel (Gjorevski et al., 2016). To date, investigation into generating skin in vitro have focused on inducing pure populations of skin cell types (e.g. dermal fibroblasts or keratinocytes) from mouse and human PSCs (Coraux et al., 2003, Ehama et al., 2007, Guenou et al., 2009, Itoh et al., 2011, Itoh et al., 2013, Mavilio et al., 2006). The logic of these approaches is that the individual parts can be combined to generate full-thickness skin (Itoh et al., 2013). This strategy, however, may lack the necessary crosstalk between cell layers to produce appendages that the current organoid approach preserves. Moreover, the organoid system may allow for induction of specific skin types (e.g., glabrous) with varying appendages. Based on our analysis, it is difficult to pinpoint the precise anatomical location represented by skin organoids; however, our data suggest that organoid derived skin may be similar to skin in the aural region—ear canal, auricular, or surrounding skin tissue. This hypothesis is supported by co-development of inner ear organoids and skin organoids, as well as expression of posterior surface epithelial and neuroepithelial marker PAX8 during differentiation (Ohyama et al., 2007). Lastly, it is notable that the skin organoids assume a cyst conformation with radially oriented HFs. Seminal work in the 1940s and 1950s described the self-assembly of skin-organoid-like structures with protruding pigmented HFs (Hardy, 1949, Moscona and Moscona, 1952). In that work, epidermal and dermal cells from mouse embryos were excised, dissociated, and re-aggregated in a hanging drop of plasma. Likewise, HF-bearing cysts arise when dissociated embryonic skin from various mammals, including humans, is subcutaneously transplanted in mice (Zheng et al., 2005, Zheng et al., 2010). More recently, a system for generating integumentary tissues was described in which mESC aggregates were partially differentiated and then placed in the kidney capsule of athymic mice (Takagi et al., 2016). Following transplantation, the aggregates preferentially differentiated into full-thickness skin layers in a cyst-like conformation and produced HFs. These results suggest that the derivation of multiple germ layers within a single stem cell aggregate may be a general strategy to produce hair-bearing skin. However, to date, this approach remains poorly defined and reliant on unknown microenvironmental factors within blood plasma or the subcutaneous and kidney capsule niches. Our study demonstrates how these skin organoid structures can be generated de novo, without the use of embryonic tissue or undefined media. We anticipate that this culture system will be useful for studying minimal cellular and microenvironment requirements for HF induction, evaluating HF growth/inhibitory drugs, or modeling skin diseases.

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