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Originally published online as doi:10.1189/jlb.0106015 on July 24, 2006

Published online before print July 24, 2006
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(Journal of Leukocyte Biology. 2006;80:816-826.)
© 2006 by Society for Leukocyte Biology

Characterization, isolation, and differentiation of murine skin cells expressing hematopoietic stem cell markers

Simone Meindl*, Uwe Schmidt{dagger}, Christine Vaculik* and Adelheid Elbe-Bürger*,1

* Department of Dermatology, Division of Immunology, Allergy and Infectious Diseases, and
{dagger} Institute of Immunology, Vienna Competence Center, Medical University of Vienna, Vienna, Austria

1 Correspondence: Department of Dermatology, Division of Immunology, Allergy and Infectious Diseases, Vienna Competence Center, Medical University of Vienna, Lazarettgasse 19, Vienna 1090 , Austria. E-mail: adelheid.elbe-buerger{at}meduniwien.ac.at


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ABSTRACT
 
As the phenotype of adult dermal stem cells is still elusive, and the hematopoietic stem cell is one of the best-characterized stem cells in the body, we tested dermal cell suspensions, sections, and wholemounts in newborn and adult mice for hematopoietic stem cell marker expression. Phenotypic analysis revealed that a small population of CD45+ cells and a large population of CD45– cells expressed CD34, CD117, and stem cell antigen-1 molecules. When cultivated in selected media supplemented with hematopoietic cytokines, total dermal cells, lineage–, and/or highly enriched phenotypically defined cell subsets produced hematopoietic and nonhematopoietic colonies. When injected into lethally irradiated recipient mice, a small percentage of newborn dermal cells was able to migrate into hematopoietic tissues and the skin and survived through the 11-month monitoring period. Our ability to isolate a candidate autologous stem cell pool will make these cells ideal vehicles for genetic manipulation and gene therapy.

Key Words: dermis • mouse • single cell suspension • semisolid medium • hematopoietic cytokines


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INTRODUCTION
 
Stem cells are responsible for the generation of all tissue types from the beginning of development up to adulthood. Evidence is growing that adult tissue stem cells might possess greater plasticity than previously thought and thus, may represent a realistic, therapeutic alternative to embryonic stem cells, yielding hope in the fields of regenerative tissue and organ repair, autotransplantations, and gene therapy applications [1 ]. The ideal stem/precursor cell population would be one that could be derived in an autologous manner from small amounts of accessible tissue biopsies. Thus, the description of stem/precursor cells with a wide spectrum of differentiation activities (adipogenic, myogenic, osteogenic, chondrogenic, neurogenic, and hematopoietic lineages) in the highly accessible tissue source, mammalian dermis [2 3 4 5 6 7 8 9 10 11 12 ], has generated much excitement in the search for alternative sources of stem and precursor cells. However, the dermal stem/precursor cell has never been phenotypically defined before differentiation.

The maintenance of stem cells requires a specific microenvironment, the niche that protects the ability of the stem cell to self-renew and prevents differentiation [13 ]. For instance, Notch signaling promotes the maintenance and self-renewal of hematopoietic stem cells (HSC) by inhibiting differentiation [14 ]. Other mechanisms through which niches regulate the fate of stem cells are by the secretion of Wnt proteins and soluble Frizzled receptors, by the production of members of the TGF-ß/bone morphogenic protein family, and by the production of Kit ligand [1 ]. Recently, the dermal papillae of hair and whisker follicles have been described as niches for endogenous, multipotent skin stem cells/precursors [4 , 6 , 8 , 15 , 16 ]. Although isolation of stem cells from hair papillae for potential therapeutic use is accessible by noninvasive techniques and free of contaminating blood, it is not practical. As the whole dermis is much easier to access, we decided to establish methods to separate defined populations of skin stem and precursor cells from this source. We investigated these cells in the newborn and adult murine dermis using hematopoietic stem cell markers, established a purification technique to reproducibly isolate dermal subpopulations, determined their colony-forming capacity, and characterized their in vivo activity.


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MATERIALS AND METHODS
 
Mice
C3H/HeN, C57BL/6 (CD45.1; Charles River Wiga GmbH, Sulzfeld, Germany, and Centre de Developement des Techniques Avanceés pour l’ Éxpérimentation Animale, Orléans, France), and C57BL/6 ROSA-26 (CD45.2; The Jackson Laboratory, Bar Harbor, ME) [17 ] mice were bred and maintained at the local animal facility. Newborn mice were obtained from timed mating, taking the appearance of a vaginal plug as Day 0 of gestation. In vitro data presented in this study are derived from C3H/HeN mice, but similar results were obtained from C57BL/6 mice. All mouse procedures were performed with consent from the Institutional Committee of Animal Experimentation of the Medical University of Vienna (Austria) and by the Austrian Ministry of Science and Education (GZ 66.009/177-Pr/4/2002; GZ 66.009/81-BrGT/2004).

Media, hybridoma, and antibodies
Normal cell culture medium consisted of RPMI-1640 medium supplemented with 10% heat-inactivated FCS (PAA Laboratories GmbH, Linz, Austria), 25 mM HEPES, 10 µg/ml gentamycin, 2 mM L-glutamine, 0.1 mM nonessential amino acids, 1 mM sodium pyruvate, 50 µM 2-ME, and 0.002% antibiotic-antimycotic solution (all from Gibco Life Technologies, Inc., Grand Island, NY). The following unlabeled FITC-, PE-, or allophycocyanin (APC)-conjugated mAb were used: anti-mouse CD3 (145-2C11, 17A2), CD4 (RM4-4), CD5/Ly-1 (53-7.3), CD11b (M1/70), CD13 (R3-242), CD19 (1D3), CD34 (RAM34), CD45 (30-F11), CD45RB (23G2), CD45R/B220 (RA3-6B2), CD62L/L-selectin (MEL-14), CD117/c-kit (2B8, ACK45), CD135/Flk-2/Flt3 (A2F10.1), stem cell antigen, Sca-1/Ly-6A/E (D7, E13-161.7), Gr-1/Ly-6C/G (RB6-8C5), and Fc{epsilon}RI (C38-2, all from BD PharMingen, San Diego, CA); CD123 (5B11, eBioscience, San Diego, CA); F4/80 (CI:A3-1, Caltag Laboratories, Burlingame, CA); and K14 (155L, Covance, Princeton, NJ). Hybridomas M5/114.15.2 (anti-I-Ab,d,q and I-Ed,k), RA3-3A1/6.1 (anti-CD45R/B220), and F4/80 were obtained from American Type Culture Collection (Manassas, VA). The hybridoma TER-119 was kindly provided by Dr. Tatsuo Kina (Department of Immunology, Institute for Frontier Medical Sciences, Kyoto University, Japan). Second-step reagents were polyclonal FITC-labeled F(ab')2 goat anti-rat IgG (H+L; Immunotech, Marseille, France), FITC-conjugated goat anti-hamster IgG (H+L; Caltag, South San Francisco, CA), streptavidin-(SAv)-PE (BD PharMingen), and SAv Texas Red (Amersham Pharmacia Biotech, Vienna, Austria). Irrelevant, isotype-matched mAb and polyclonal antibodies were used as negative controls. Some of the mAb used were purified from supernatants of the corresponding hybridomas, and protein concentrations were adjusted to 1 mg/ml before they were FITC-conjugated or biotinylated.

Single cell suspension of dermal cells
In addition to adult skin, we used newborn skin to enhance the possibility of isolating cells more enriched for multipotent or even pluripotent stem cell activity, as dermal tissue is rapidly expanding at this stage of development. In addition, it has been shown that skin-derived precursors are more abundant in neonatal than adult skin [8 ]. Potential stem cell contamination sources in the newborn and adult dermis (e.g., muscle, adipose tissue) were removed. We minimized the risk of blood contamination by careful washing of the remaining dermis in PBS. Dermal cell suspensions from whole body skin were prepared essentially as described [18 ]. Briefly, body skin from adult mice was shaved, sterilized (betaisodona/PBS and rinsed with 70% ethanol), and placed dermal-side down on 0.8% trypsin/PBS (45 min, 37°C). After separating epidermis and dermis, tissues were vigorously agitated in a shaking water bath (40 min, 37°C) in FCS-supplemented RPMI 1640 containing 0.025% DNase. After filtering the dermal cell suspension, viability was determined by trypan blue exclusion.

Analysis and purification of dermal cells and their subpopulations
For the evaluation of Lin+ cells in the newborn and adult dermis, dermal cell suspensions were labeled with mAb directed against lineage (Lin) markers [dendritic cells (DC): anti-MHC Class II (M5/114); macrophages: anti-F4/80; monocytes/macrophages: anti-CD11b (M1/70); plasmacytoid DC/B cells: anti-CD45R/B220 (RA3-6B2); T cells: anti-CD3 (17A2); granulocytes/monocytes/plasmacytoid DC: anti-Gr-1 (RB6-8C5); and erythrocytes: anti-Ly-76 (TER-119)] and analyzed by flow cytometry. The number of Lin+ cells was expressed as the percentage of total viable leukocytes. Depletion of Lin+ cells was achieved by labeling dermal cells with a cocktail of mAb (30 min, 4°C) described above, followed by low-tox-M rabbit complement (45 min, 37°C, Cedarlane Laboratories, Hornby, Ontario, Canada) in a shaking water bath (negative selection). Dead cells together with large cells such as fibroblasts were removed by density gradient centrifugation (Lympholyte M, Cedarlane Laboratories). All cells removed by this procedure are termed as Lin+ cells. For certain experiments, total dermal cells or Lin– cell populations were sorted for high purity using a mismatched panning technique [19 ]. Cells were labeled with rat anti-mouse mAb directed against Sca-1 or CD45 (30 min, 4°C). After extensive washes, cells were transferred into petri dishes coated with goat anti-mouse Ig (60 min, room temperature) and incubated (30–40 min, 37°C). Nonadherent cells were gently rinsed off using prewarmed PBS. Adherent cells were released by adding excess amounts of mouse {gamma}-globulin (15 min, 4°C). The purity of isolated cells was assessed by flow cytometry. These cells were used for further in vitro and in vivo experiments.

Preparation of bone marrow (BM)
Six- to 8-week-old mice were killed, and their femurs and tibias were removed aseptically. Marrow cavities were flushed with medium containing 0.025% DNase using a syringe fitted with a 22- or 24-gauge needle. Single cell suspensions were prepared by repeated pipetting, and particulate matter was removed by passing the cells through a nylon mesh. Red blood cell lysis was performed using ammonium chloride (0.8% NH4Cl/0.1 mM EDTA). Mononucleated cells were washed two times in standard medium and counted using a hemocytometer.

Clonogenic assay
The colony-forming potential of total dermal cells or purified subpopulations was examined using colony-forming assays. Dermal cells were plated in duplicates into petri dishes containing MethoCult GF M3434 [1% methylcellulose in IMDM, 15% FBS, 1% BSA, 10–4 M 2-ME, 2 mM L-glutamine, 10 µg/ml recombinant human insulin, 200 µg/ml human transferrin, 50 ng/ml recombinant mouse stem cell factor (SCF), 10 ng/ml recombinant mouse IL-3, 10 ng/ml recombinant human IL-6, and 3 units/ml recombinant human erythropoietin (Stem Cell Technologies, Vancouver, Canada)] for selected time periods. To test cytokine specificity, cells were simultaneously cultured in MethoCult GF M3231 (1% methylcellulose in IMDM, 30% FBS, 1% BSA, 10–4 M 2-ME, 2 mM L-glutamine, Stem Cell Technologies). Mononucleated cells from peripheral blood and BM samples were processed as above and used as controls. After incubation for 7, 10, and 21 days at 37°C and 5% CO2 in a humidified atmosphere, colonies (colony≥8 cells) were scored based on morphologic characteristics, and individual colonies were picked arbitrarily under the microscope. Cytospin preparations, histochemistry staining, and flow cytometry analysis of these cells were performed as described below.

Liquid culture
In certain experiments, Lin– dermal cells from newborn mice were cultured (1x106 cells/ml) in Stem-Pro-34 medium including nutrient supplement (kindly provided by Dr. Hartmut Beug, IMP, Vienna, Austria) and 100 ng/ml mouse recombinant SCF and 2 ng/ml recombinant murine IL-3 (PeproTech Ltd., London, UK) at 37°C and 5% CO2 using four-well culture plates (Costar, Roskilde, Denmark). After 24 h, nonadherent cells were transferred into new wells. Partial medium changes were performed daily, and cell numbers were determined starting at Day 5 of culture using an electronic cell counter (CASY, Schärfe-System, Reutlingen, Germany). At Day 35 of culture, cells were cytocentrifuged and stained with the mast cell-specific dye toluidine blue (Sigma-Aldrich, St. Louis, MO) and analyzed for the expression of selected surface markers by FACS analysis.

Histochemistry
Petri dishes were scored for cell growth at different time-points. Individual colonies were cytocentrifuged onto glass slides and air-dried (30 min, room temperature) before further treatment. Cells were determined with the following staining techniques. Adipocytes and sebocytes were identified by the cellular accumulation of neutral cytoplasmic lipid vacuoles, which can be visualized with oil red O (Sigma-Aldrich). Cells were cytocentrifuged and fixed in 3.7% formaldehyde for 5 min. Fixed cells were rinsed in distilled water and incubated with oil red O (60% in distilled water) for 10 min at room temperature. Staining background was cleared using 60% 2-propanol for 30 s.

Cell morphology was determined by May-Grünwald-Giemsa staining (Merck, Darmstadt, Germany). Cells were rinsed in PBS, cytocentrifuged, and fixed in methanol for 5 min, washed two times in PBS, and stained with May-Grünwald’s eosin methylene blue (3 min, room temperature), followed by two washing steps in PBS. For counterstaining with Giemsa, cells were incubated in Giemsa’s azure eosin methylene blue working solution (20 min, room temperature, Merck), washed in tap water, and air-dried.

Mast cells were identified by toluidine blue staining. Cells were incubated (10 min, room temperature) in toluidine blue (10% in methanol, Sigma-Aldrich). Afterward, cells were washed in tap water and air-dried. Images were collected using an inverted microscope with a Nikon Coolpix 995 digital camera (Nikon, Tokyo, Japan).

Immunofluorescence staining and evaluation of immunofluorescence results
Skin biopsies from newborn and adult mice were embedded in OCT Tissue-Tek (Sakura Finetek Europe, Zoeterwoude, The Netherlands), snap-frozen in liquid nitrogen, and stored at –80°C until further processing. Frozen tissue was cut into 5 µm thick sections and mounted on capillary gap microscope slides (Dako, Glostrup, Denmark). Sections were air-dried for 20 min, fixed in ice-cold acetone for 10 min, and washed three times in PBS. In certain experiments, dermal wholemounts of mouse skin were prepared. Skin was cut into pieces (0.5x0.5 cm2) and placed dermal-side down on 3.8% ammonium thiocyanate/PBS (25 min, 37°C). After removal of the epidermis, the dermis was washed in PBS, fixed in acetone (10 min, room temperature), and rehydrated in PBS. Prior to the staining procedure, tissue was incubated with rat serum [5% (v/v) in PBS, 15 min, room temperature] to block unspecific binding of antibodies. To identify epidermal and dermal subpopulations, sections and wholemounts were incubated with FITC- and PE-conjugated mAb directed against selected mouse antigens. Immunostained sections and wholemounts were analyzed using a confocal laser-scanning microscope (LSM 510, Zeiss, Oberkochen, Germany).

For two- and three-color analyses, dermal cells (5x105/sample) were resuspended in cold FACS buffer (1% FCS/PBS/0.01% NaN3) and incubated with APC-, FITC-, and PE-conjugated or biotinylated antibodies directed against selected mouse antigens. Biotinylated antibodies were followed by SAv-PE, SAv-FITC, or SAv-TRC. To block nonspecific binding of antibodies, cells were incubated with mouse/goat serum [5% (v/v) in PBS, 20 min, 4°C] prior to the staining procedures. Specificity of staining was confirmed using isotype-matched control antibodies. Dead cells were excluded by 7-amino-actinomycin D (7-AAD; 1 µg/ml, Sigma-Aldrich) uptake. In some experiments, only cells with a profile described for lymphocytes were gated, whereas in others, the whole viable population was analyzed. Fluorescence was measured using a FACScan or FACSCalibur flow cytometer, and data were analyzed with CellQuest software (Becton Dickinson, Mounting View, CA). To detect intracellular antigens, dermal cells were permeabilized with Cytofix/Cytoperm (250 µl/ml, 20 min, 4°C, Becton Dickinson). After washing with 0.1% saponin/PBS, cells were stained with nonconjugated or direct-labeled antibodies (30 min, 4°C). Nonconjugated antibodies were visualized by a second-step reagent (30 min, 4°C). Appropriate isotype antibodies were used as controls. After washing with 0.1% saponin/PBS, subsequent staining was performed as described above.

Long-term competitive reconstitution assays
To distinguish between host and donor cells, we used C57BL/6 ROSA-26 (CD45.2) mice as donors, which express ß-galactosidase (ß-gal) systemically. Consequently, these cells can be identified by X-gal staining. Donor cells were injected into congenic mice expressing the CD45.1 allele, allowing analysis of donor and host cells for defined populations. Adult (8- to 12-week-old) female C57BL/6 (CD45.1) mice were given sterile, acidified water (pH 3.0) ad libitum for 2 weeks. Subsequently, recipient mice were exposed to 7.5 Gray total body {gamma}-irradiation in a split dose with a 3-h interval. Five hours after irradiation, inocula containing total dermal cells (1x107/mouse), Lin– dermal cells (1x107/mouse), or CD45+ dermal cells (1x103–1x104/mouse) from C57BL/6 ROSA-26 newborn mice or for control purposes, BM cells (1x107/mouse) from C57BL/6 ROSA-26 adult mice were injected i.v. into the tail vein or i.p., using a syringe fitted with a 25-gauge needle in the presence or absence of syngeneic BM (1:10). Control mice received 50 µl PBS. At least three mice per group were used for each experimental condition. Total dermal cells from adult but to a lesser extent, from newborn mice often failed to reconstitute i.v.-transplanted recipient mice, because of cellular aggregation causing fatal embolic formation. To determine donor chimerism, tail-vein blood samples were first analyzed 6 weeks after transplantation and then every second month. In addition, recipient mice were killed at selected time-points (8–44 weeks), and cells from BM, spleen, liver, lymph nodes, thymus, epidermis, and dermis were double-stained with anti-CD45.2 and a panel of mAb against Lin markers to monitor engraftment of donor-derived cells.

ß-Gal detection
X-Gal staining of cell suspensions isolated from selected organs (BM, spleen, liver, lymph nodes, thymus, epidermis, and dermis) was performed as described [20 ]. Air-dried cytospin preparations were fixed with 0.5% glutaraldehyde (15 min, room temperature, Sigma-Aldrich) and washed three times with PBS. Specimens were then incubated overnight with the ß-gal substrate, 5-bromo-4-chloro-3-indolyl-ß-D-gal (X-gal, Sigma-Aldrich) in a humidified chamber at 37°C and subsequently, washed in tap water.


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RESULTS
 
Hematopoietic and nonhematopoietic dermal cells express stem cell markers
Freshly isolated, total dermal cells from newborn (3.5x106 cells/mouse) and adult (5.5x106 cells/mouse) mice were analyzed by flow cytometry for the presence of hematopoietic cell surface markers (Table 1 ). In repeated experiments, there was an approximately fivefold greater yield of Lin+ (Table 2 ) and CD45+ cells (Fig. 1A ) in the adult dermis compared with newborn dermis. With regard to CD45 expression, we found two discrete populations in the adult dermis. CD45high cells coexpressed CD3, representing T cells (data not shown). A small population of CD45+ cells from newborn and adult dermis displayed markers associated with early progenitors and stem cells such as Sca-1, CD34, and CD117 (Fig. 1A) [21 22 23 ]. Although newborn and adult dermis contained CD45+Sca-1low cells, only adult dermis harbored CD45+Sca-1high cells as determined by flow cytometry (Fig. 1A , rectangles), the latter of which were also detectable in skin sections (Fig. 1B) , reflecting the superior sensitivity of the flow cytometry method. Furthermore, the newborn dermis but not the adult dermis contained small populations of CD45+CD34+ and CD45+CD117+ cells. Although 85% of CD45+Sca-1+ in newborn mice coexpressed CD117, the percentage of CD45+Sca-1+CD117+ cells decreased dramatically with age (5% in adults; Fig. 1A , histograms). This observation argues in favor of a role for HSC in early postnatal development. It is interesting that ~11% of CD45– newborn dermal cells expressed Sca-1. As compared with the newborn dermis, a greater number (54%) of adult dermal cells were positive for Sca-1. In this regard, a similar expression profile for Sca-1+ cells has been described most recently in neonatal and adult murine hearts [24 ].


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  		Table 1. Cellular Recoveries of Dermal Cells from Newborn and Adult Mice


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  		Table 2. Frequencies of Lin+ Cells in the Newborn and Adult Dermis


Figure 1
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  		Figure 1. Dermal cells in mice express hematopoietic stem cell markers. (A) Total dermal cells from newborn and adult mice were stained for the indicated markers and analyzed by flow cytometry. Quadrants were set based on isotype control staining (not shown), and numeric values indicate percentage among mononuclear cells. To better characterize dermal cells for stem cell markers, triple staining was performed with FITC-, PE-, and APC-conjugated mAb. The rectangles represent the gate for detection of the third antibody analyzed in the histograms. Dead cells positive for 7-AAD were excluded, and 50,000 (dot plots) or 100,000 (histograms) events per sample were acquired. Whereas open profiles in the histograms illustrate the reactivity of Sca-1+CD45+ cells with an anti-CD117 mAb, closed profiles represent an irrelevant, isotype-matched control mAb. These results are representative of those obtained in five independent experiments. (B) Dermal wholemounts from newborn and adult mice were exposed to FITC-anti-Sca-1/PE-anti-CD45 double-labeling. Arrows denote double-positive cells. Micrographs were taken by confocal laser-scanning microscopy; original scale bars: 20 µm.

Newborn dermal cells possess greater clonogenic potential than adult dermal cells
To assess the clonogenic potential of newborn and adult dermal cells, they were seeded at selected cell numbers into petri dishes with methylcellulose supplemented with cytokines promoting hematopoietic differentiation. BM cells were used as positive controls. Colonies were observed consistently after 7 days of culture (data not shown) and increased in number and size during the following days. After 21 days, three distinct populations of cells were routinely observed under these conditions: 1) cells with lipid vacuoles, 2) small, piled cells (the center of the colony appears light-brown), and 3) shiny, "white" cells with a clear cytoplasm (Fig. 2A , photomicrographs). The generation of the latter two cell populations was strictly dependent on the presence of exogenous factors (data not shown). Although a degree of variation was observed between experiments with respect to the number of colonies per seeded cells, we found that the colony-forming capacity was always ≥4-fold greater in newborn relative to adult dermal cells (Fig. 2A , graphs). Flow cytometry analysis revealed that development from newborn and adult cultured dermal cells gave rise to 72% and 4% of hematopoietic cells, expressing the pan-leukocyte marker CD45, respectively (Fig. 2B) . Populations with bright and dim CD45 expression were observed. Staining of individual colonies revealed that CD45bright cells expressed markers described previously for the identification of mature mast cells such as Fc{epsilon}RI, Sca-1, CD13, CD34, and CD117 (Fig. 2C) [25 ]. In addition, these colonies exhibited morphologic features of mast cells such as inclusion of metachromatic granules visualized by toluidine blue staining (data not shown). CD45dim cells represented white colonies and expressed MHC Class II, F4/80, Gr-1, and CD123, whereas CD11b, CD34, and CD45R/B220 were absent (Fig. 2D and data not shown), suggesting a differentiation of dermal stem/precursor cells to the myeloid-macrophage pathway. Oil red O staining of intracellular lipids in cells isolated from individual colonies revealed their adipocyte and/or sebocyte nature (data not shown). Using the identical assay, peripheral blood samples collected from newborn and adult mice failed to generate colonies.


Figure 2
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  		Figure 2. Dermal cells from newborn mice have a greater clonogenic potential compared with adult mice. (A) Freshly isolated, total dermal cells from newborn and adult mice were plated at the indicated densities into methylcellulose (MethoCult GF M3434) as described in Materials and Methods. After 21 days, different types of colonies could be observed (upper panels). The lower panels show the frequencies of colonies per seeded cells. Photomicrographs are representative of eight (newborn) and six (adult) independent experiments. (B) All colonies were harvested on Day 21 and stained with a PE-labeled, anti-CD45 mAb. Dead cells positive for 7-AAD were excluded, and 10,000 events per sample were acquired. (C) Individual colonies with mast cell morphology were picked, pooled, and stained for the indicated markers and analyzed by flow cytometry. Three thousand events per sample were acquired. (D) FACS analysis of individual colonies with a white phenotype. Histograms show 2000 (Exp. 1) and 1000 (Exp. 2) events per sample. Open profiles illustrate the reactivity with the relevant mAb; closed profiles represent irrelevant, isotype-matched control mAb.

Precursors of lipid+ dermal cells express Sca-1
As we have removed the lower dermis and the panniculus adiposus, both containing adipocytes, and have not used an adipogenic medium to differentiate lipid+ cells, it is conceivable that the majority, if not all, oil red O+ cells represent sebocytes, implying that our dermal cell suspensions may be "contaminated" with epidermal cells. To test this hypothesis, freshly isolated dermal cell suspensions were analyzed by flow cytometry for the expression of K14 and Sca-1. It was surprising that the majority of dermal cells (89% in newborn, 69% in adult mice) were positive for K14, and 14% (newborn) and 37% (adult) thereof coexpressed Sca-1 (Fig. 3A , dot plots). The anatomical location of these cell populations was assessed in skin sections and dermal wholemounts. As expected, K14 was expressed in the basal layer of the interfollicular epidermis, the outer root sheath of hair follicles, and the sebaceous gland (Fig. 3A and data not shown). Sca-1+K14+ cells were localized in the follicle papillae in adult skin but could not be detected in newborn skin, most likely as a result of the low-level expression of these molecules (Fig. 3A , lower panel). Furthermore, we found that Sca-1+K14– cells colocalized with the endothelial marker CD31 (Fig. 3A , lower panel). As Sca-1 is not only a hematopoietic stem cell marker but can also be found in nonhematopoietic tissues such as the vasculature of several organs [26 27 28 29 ], our data strongly suggest that Sca-1 identifies vascular endothelial cells/precursors in the skin, which is in line with our assumption that some Sca-1+ cells have a mesenchymal cell nature.


Figure 3
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  		Figure 3. Sca-1+ skin cells colocalize with K14 and CD31. (A, B) Total and Lin– dermal cell suspensions from newborn and adult mice were stained for the indicated markers and analyzed by flow cytometry. Regions were set based on isotype control staining (not shown), and numeric values in the table indicate percentage among mononuclear cells in the regions (+/–: dim; +: intermediate; ++: bright). Skin sections and wholemounts from newborn and adult mice were exposed to FITC-anti-K14 single-labeling and PE-anti-Sca-1/FITC-anti-K14 and FITC-anti-Sca-1/PE-anti-CD31 double-labeling. Micrographs were taken by confocal laser-scanning microscopy; original scale bars: 20 µm.

After Lin depletion by negative selection with antibody and complement treatment, the total amount of Sca-1+ cells enriched twofold in newborn but not in adult dermal cells (Fig. 3B) . Furthermore, depletion of Lin+ cells resulted in a moderate (one- to twofold) enrichment (14% vs. 30% in total vs. depleted newborn dermis; 37% vs. 44% in total vs. depleted adult dermis) of K14+Sca-1+ cells (Fig. 3B) . The cellular recoveries from newborn and adult dermal cells per mouse were 47% and 17%, respectively (Table 1) . By testing the clonogenic potential of Lin– newborn and adult dermal cells, we found by morphologic and phenotypic criteria that after 21 days of culture, the majority of the colonies contained cells with lipid vacuoles (Fig. 4A , photomicrographs), which were visualized by oil red O staining (data not shown). By flow cytometry, approximately one-third of the cells derived from newborn and adult Lin– dermal cells expressed K14, and 1–2% displayed anti-CD45 reactivity (Fig. 4A , dot plots). Whether the CD45–K14– cells are differentiated sebocytes that have lost K14 expression or represent adipocytes remains to be determined.


Figure 4
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  		Figure 4. The majority of Lin–Sca-1+ dermal cells coexpress K14. (A) Lin– dermal cells (1x105/well) were plated into methylcellulose (MethoCult GF M3434). At Day 21, photomicrographs of outgrowing colonies were taken. Subsequently, all cells were harvested and stained for CD45 and K14. Dead cells positive for 7-AAD were excluded, and 10,000 (newborn) and 15,000 (adult) events per sample were acquired. Data are representative of three independent experiments. (B) Lin– dermal cells from newborn and adult mice were purified (histograms: open profiles illustrate the reactivity with the relevant mAb; closed profiles represent irrelevant, isotype-matched, control mAb) for Sca-1+ cells by positive selection as described in Materials and Methods, stained for the indicated surface and cytoplasmic markers, and analyzed by flow cytometry. Dead cells positive for 7-AAD were excluded, and 5000 events per sample were acquired.

To investigate whether Sca-1+ cells are indeed precursors of lipid+ cells, Lin– dermal cell suspensions were enriched for Sca-1+ cells to high purity (Fig. 4B) . Flow cytometry analysis revealed that most (89%) Lin–Sca-1+ cells from newborn mice coexpressed CD34, only a few displayed CD45 reactivity, and all were negative for CD117. Lin–Sca-1+ adult dermal cells failed to express CD34 and CD117, but a considerable percentage was positive for CD45. A great percentage of newborn and most of adult Lin–Sca-1+ dermal cells expressed K14 at moderate and high levels (Fig. 4B) . Lin–Sca-1+ newborn dermal cells contained Sca-1brightK14– and Sca-1brightK14int cell populations as opposed to adult dermal cells. By testing the clonogenic potential of newborn Lin–Sca-1+ cells, we found by morphologic and phenotypic criteria solely colonies with lipid+ cells after 21 days (data not shown). In cultures with adult Lin–Sca-1+ cells, significantly fewer lipid+ cells were present compared with newborns, and in addition, other types of colonies could be observed (data not shown). Taken together, these results lead us to conclude that the anti-Sca-1 mAb can be used to identify a precursor of lipid+ cells in the skin.

Generation of large numbers of mast cells from Lin– dermal cells in liquid culture
Semisolid culture of total dermal cells with hematopoietic growth factors resulted mainly in the outgrowth of mast cells. To test whether mast cells can be generated in a serum-free liquid culture system, we used the mast cell proliferation factors SCF and IL-3 and Lin– dermal cells from newborn mice as a starting population. After selected time-points, the cultured cells were characterized by FACS and histological analysis. With both cytokines present, in each of two independent experiments, we observed an expansion of cells for approximately 5 weeks. We generated 2 x 109 mast cells from a starting population of 1 x 106 Lin– dermal cells/ml (2000-fold increase), identified as toluidine blue+ and CD45+Fc{epsilon}RI+Sca-1+CD34+CD117+ cells by 35 days of culture, and no outgrowth of cells was observed in the absence of SCF and IL-3 (Fig. 5A 5C, and 5D ). Similar cell numbers could be obtained when mast cells were generated in the presence of SCF and IL-3 using total BM (0.5x106 cells/ml) as a starting source (data not shown). All dermal-derived cells contained metachromatic granules (Fig. 5C) and failed to express surface markers for early hematopoietic progenitor cells (CD135), granulocytes/monocytes (Ly-6G/Ly-6C, CD62L), DC (MHC Class II), T cells (CD3, CD4, CD5, CD62L, CD90), or B cells (CD5, CD19, CD62L; data not shown). After the initial phase of rapid expansion, the proliferation rate gradually decreased, and the cells stopped to proliferate after approximately 5 weeks of culture. When proliferating, dermal-derived cells were transferred at Day 12 into methylcellulose containing hematopoietic differentiation factors, two types of colonies were detected 16 days later: 1) mast cell colonies (65.7%) and 2) white colonies (35.3%; Fig. 5B and data not shown). These data corroborate and expand a report by Yamada et al. [30 ], which identified CD45+CD117+Fc{epsilon}RI– immature mast cells in fetal murine skin, which could be differentiated into mature mast cells in the presence of SCF and FCS.


Figure 5
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  		Figure 5. Enormous numbers of mast cells can be generated from Lin– dermal cells in liquid culture. (A) Lin– newborn dermal cells (1x106/ml) were cultured in the presence or absence of SCF and IL-3 as described in Materials and Methods. Each rectangle in the graph corresponds to the cumulative cell number calculated on the indicated day of culture. (B) After 12 days in liquid culture, cells were harvested, washed, and plated (1x105/ml) in methylcellulose (MethoCult GF M3434). Photomicrographs show colonies after 16 days in methylcellulose. (C) After 34 days in liquid culture, cells were cytocentrifuged and stained with toluidine blue. (D) The same samples were labeled for flow cytometry analysis, and the x- and y-axis of the histograms represent fluorescence intensity and cell numbers, respectively. Whereas open profiles illustrate the reactivity with the relevant mAb, the closed profile represents irrelevant, isotype-matched, control mAb. Dead cells were excluded, and 50,000 events per sample were acquired. Data shown are representative of two independent experiments. Original scale bar: 20 µm.

The majority of Lin–CD45+ dermal cells is mast cell precursors
Our observation that hematopoietic activity still occurred after Lin depletion prompted us to further sort Lin– dermal cells for CD45-reactive cells. After positive selection with an anti-CD45 mAb, we regularly obtained 3.7 x 103 (range: 4x102–8.3x103) and 1.8 x 104 (range: 1.6x103–1.4x104) CD45+ cells, representing a cellular recovery of 0.01% and 0.03% per newborn and adult mouse, respectively (Table 1) . The viability of CD45+ cells ranged from 75% to 90% and had a reproducible purity of ≥89% (Fig. 6A ). To characterize these cells, one part was stained and analyzed by flow cytometry, and the other part was seeded into methylcellulose, supplemented with cytokines providing hematopoietic differentiation to assess their clonogenic potential. Eighteen percent and 25% of the freshly isolated CD45+ cells from newborn as well as adult mice, respectively, were positive for Sca-1. All freshly isolated Sca-1+CD45+ cells from newborn mice coexpressed CD117 and CD34 (Fig. 6A , dot plots). After 21 days of culture, plating of newborn Lin–CD45+ cells in semisolid medium revealed that these cells mainly (~90%) formed mast cell colonies, as determined by May-Grünwald-Giemsa staining (Fig. 6B) . Occasionally, another cell type with large nuclei, little cytoplasm, and no metachromatic granules could be detected. Similar results were obtained when we cultured Lin–CD45+ dermal cells isolated from adult mice, although the numbers of the colonies per seeded cells were less than 70% compared with newborn cells (data not shown). Lipid-containing cells were observed sporadically in certain experiments at low numbers (data not shown). When plating the Lin–CD45– cell fraction, we occasionally observed dividing cells and colonies with cells containing lipid vacuoles.


Figure 6
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  		Figure 6. Lin–CD45+ dermal cells differentiate into mast cells in semisolid media. Lin– dermal cells were purified for CD45+ cells by positive selection as described in Materials and Methods (histograms: open profiles illustrate the reactivity with the relevant mAb; closed profiles represent irrelevant, isotype-matched, control mAb). One part of the cells was stained for surface marker expression and analyzed by flow cytometry. Dead cells positive for 7-AAD were excluded, and 5000 events per sample were acquired. Remaining cells from newborn mice were plated in methylcellulose (MethoCult, GF M3434). At Day 21, cells were harvested, and their morphology was assessed by May-Grünwald-Giemsa staining (B). The black arrow displays a differentiated mast cell with granules, and the white arrow marks a cell with a high nucleus to cytoplasm ratio. Data are representative of three similar experiments. Original scale bar: 20 µm.

Dermal cells engraft into hematopoietic tissues and the skin in vivo
Although results from our clonogenic assays strongly suggested the presence of hematopoietic potential, this in vitro assay detects hematopoietic progenitors, which include but are not exclusively HSC. To determine the presence of HSC activity in these cultures, we assessed their capacity to generate long-term, multilineage hematopoietic reconstitution of myeloablated recipient animals. For this, we used a competitive repopulation assay [31 ], in which a given cell population is required to compete in the same recipient with a genetically distinguishable standard source of HSC activity. Total dermal cells, Lin– and CD45+ dermal subfractions isolated from C57BL/6 ROSA-26 (CD45.2) newborn mice, and for control purposes, BM from adult C57BL/6 ROSA-26 mice were transplanted into lethally irradiated, congenic C57BL/6 (CD45.1) mice (Fig. 7A ). Recipient mice injected i.p. or i.v. with dermal cells or PBS alone demonstrated 100% mortality within 10 days. Although mice receiving dermal cells and syngeneic BM cells i.p. also died within 10 days, mice injected i.v. with dermal cells and syngeneic BM survived up to 44 weeks. Eleven months following transplantation, the appearances of the spleen, liver, lymph nodes, and thymus were normal among the different groups. No hematopoietic nodules on the surface of the spleen could be observed. These examinations indicated that the homing potential of dermal cells is not altered compared with BM cells. Randomly selected mice from each group were scanned for the presence of donor cells at various time-points. During investigation, no donor cells were identified by flow cytometry analysis in the peripheral blood. However, donor-derived dermal cells (CD45.2+, ß-gal+) were sporadically but repeatedly detected in recipient hematopoietic tissues (BM, spleen, liver, lymph nodes, thymus) and the skin (data not shown) between 2 and 11 months post-transplantation as determined by flow cytometry analysis and X-gal staining (Fig. 7B and 7C) . No obvious differences in the repopulation capacity were observed when dermal cells, Lin–, or CD45+ dermal cells were injected.


Figure 7
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  		Figure 7. Dermal-derived cells incorporate into hematopoietic tissues and the skin. (A) Experimental design. Lethally irradiated mice (CD45.1) were transplanted with total dermal cells, Lin–, and CD45+ dermal cells from newborn C57BL/6 ROSA 26 (CD45.2) mice together with syngeneic (CD45.1) BM cells. Chimerism was evaluated by flow cytometry and ß-gal detection. (B) CD45.2+ donor cells and ß-gal activity (blue cells) were observed in the BM of mice injected with purified (≥98%) CD45+ dermal cells 12 weeks after transplantation. (C) Photomicrographs show ß-gal activity in the spleen of mice reconstituted with Lin– dermal cells, 38 and 44 weeks after transplantation.


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DISCUSSION
 
In the present study, we have demonstrated that the skin contains CD45+ and CD45– populations of cells that can be distinguished by the expression of HSC markers such as Sca-1, CD34, and CD117. Moreover, we have further developed a technique, which allowed us to purify subpopulations to high purity and viability and to analyze their differentiation capacity in vitro and in vivo. Furthermore, our observations add to the growing number of antigens available for immunotyping skin endothelium and raise intriguing questions about the significance of shared gene expression patterns between endothelium and stem or progenitor cells of hematopoietic and nonhematopoietic tissues.

Our data demonstrate that the skin contains undifferentiated precursors, which can be induced in vitro to give rise to lipid+ cells. To characterize these precursors, dermal cells were stained with markers directed against Sca-1, mesenchymal, and epithelial cells. It is surprising that the dermal cell populations contained significant numbers of K14+ cells, implying an epidermal contamination. In preliminary experiments, we found that a minor population of K14+ cells in newborn (6%) and adult (8%) dermal cell suspensions expressed high levels of {alpha}6-integrin and low levels of CD71 (data not shown), a marker profile similar to that reported for epidermal stem cells [32 ]. As some of these cells also expressed Sca-1, the phenotype is somewhat compatible with the side population cells ({alpha}6-integrin+ß1-integrin+Sca-1+K14+) recently described in mouse skin [33 ]. It is interesting that the vast majority of the Sca-1+ cells in newborn but not in adult mice coexpressed CD34 and thus, are reminiscent of a Sca-1+CD34+ cell population, most recently described, representing adipocyte precursors in the hypodermis of neonatal mice [29 ]. Although we have carefully removed the lower dermis and the hypodermis, we identified these cells in dermal cell suspensions, implying that some Sca-1+CD34+ cells are also present in the upper dermis in the newborn skin. In fact, we found these cells to be more evenly distributed in newborn dermis (C. Vaculik, unpublished observation) compared with older mice, where they seem to be preferentially located in the hypodermis [29 ].

As a certain percentage of lipid+ cells express K14, it is conceivable that our cultures contain sebocytes and adipocytes, which may arise from distinct Sca-1+ precursors present in our starting population (Sca-1+K14–, Sca-1+K14+, Sca-1+K14++). Single cell experiments need to be done to address this assumption. A role for Sca-1 in adipocyte development is supported by the observations that Sca-1–/– mice exhibit a deficiency in adipocyte colony-forming cells [34 ] and that Sca-1+ cardiac precursors produce adipocytes in culture [24 ]. Further, it has been shown that adipocyte-like cells can be generated from rodent follicle dermal cells containing Sca-1+ cells using an adipogenic medium [2 , 15 , 24 ], although their adipocyte nature has not been proven.

Another remarkable observation was that a certain percentage of Lin–CD45+ dermal cells in newborn, but less so in adult, mice coexpressed Sca-1, CD117, and CD34 and did not contain metachromatic granules (data not shown). Classically, CD34+CD45+ cells represent a late stage of HSC differentiation, which have become activated when they have migrated from the BM to the secondary tissues. The earliest HSC in the BM are Lin–Flt3–Sca-1+CD11b–CD34–CD45+CD90+CD117+. After this stage, they populate various tissues and down-regulate Sca-1 while up-regulating CD34 [23 , 35 36 37 38 ]. It is intriguing that a considerable proportion of freshly isolated, newborn dermal Lin– CD34+CD45+CD117+ cells was Sca-1+. Our further observation that Lin– dermal cells can differentiate into mast cells and white cells in the presence of SCF and IL-3 in semisolid medium as well as in liquid cultures suggests that different precursors are present in the dermis, which respond to SCF and IL-3 or that one common precursor exists, which can develop into different cell types. Cells with a phenotype comparable with our dermis-derived cells have been described to be present in the murine BM (Fc{epsilon}RI–Sca-1–CD13+CD34+CD117+) and circulating fetal murine blood (Lin–Fc{epsilon}RI–CD90lowCD117+) and required SCF and IL-3 to differentiate into mature mast cells [39 , 40 ]. Subsequently, Yuan et al. [41 ] demonstrated that SCF or IL-3 can promote the in vitro growth of Fc{epsilon}RI–Sca-1+CD117+ and Fc{epsilon}RI–Sca-1–CD117+ populations from the BM of adult mice as well as their full maturation into mast cells. Most recently, it has been shown that Lin–Sca-1+CD45+CD117+Fc{epsilon}RI+ cells already exhibit several features of connective tissue-type mast cells (e.g., inclusion of metachromatic granules, histamine release on ligation of surface IgE, mRNA expression for mast cell protease-1, -4, -5, and -6). These cells appear upon culture of vibrissae follicles isolated from adult mice in the presence of a SCF-containing medium [41 ]. However, in contrast to our study, the phenotype of the precursor has not been determined. These results together with the findings that hair follicles contain small numbers of CD117+ cells and produce SCF suggest that hair follicles may provide a unique microenvironment for local development of mast cells [42 43 44 ]. Whether hair follicle-derived mast cells correspond to those that we have identified needs to be investigated carefully. It also remains to be determined whether the cell types we have identified (e.g., Lin–Sca-1+CD117–K14–, Lin–Sca-1+CD117–K14+) can give rise to cells of other lineages when using appropriate differentiation media (e.g., lymphoid, osteocytic, neural lineages) or whether they are fully committed to the myeloid/adipocyte/sebocyte lineages.

We are aware that our study contains some drawbacks. It is conceivable that our observations reflect the in vitro expansion and differentiation of HSC present as blood contaminants in our dermal cell preparations. This possibility needs to be formally excluded, although no outgrowth of cells was observed when peripheral blood samples were cultured under the same conditions. Alternatively, our dermal cell preparations may contain small numbers of mature mast cells, which can re-enter the cell cycle when placed in culture. However, mature mast cells have not been detected within our freshly prepared Lin– dermal cell preparations (data not shown).

We have demonstrated long-term (11 months) engraftment of dermal cells into hematopoietic tissues and the skin in lethally irradiated mice. Our failure to detect donor cells in the peripheral blood by flow cytometry might be a result of their low frequency. In fact, we found few donor cells in BM, spleen, liver, lymph nodes, and thymus throughout the observation period of 2–11 months. Therefore, their further phenotypic analysis was unfeasible. One possible explanation for the poor reconstitution capacity of dermal cells in the tissues could be a disadvantage to these cells during a competitive repopulation assay in the presence of adult stem cells. A second possibility is that dermal-derived cells are targets of host-derived NK cells. It remains to be tested whether the treatment of mice with anti-asialo GM1 antiserum will facilitate stem cell engraftment. It is interesting that we regularly observed a small number of X-gal+ donor cells in dermal cell suspensions. It remains to be investigated whether they are distributed uniformly in the dermis or homed to particular niches such as the dermal papillae of hair, as most recently proposed [8 ]. Nonetheless, our data suggest a homing effect of dermal stem cells to their structural "niche" in the skin, analogous to that of HSC to the BM and extramedullary tissues such as the spleen, where they associate with sinusoidal endothelial cells [45 , 46 ]. A variety of recent studies imply that endothelial cells create a niche in hematopoietic tissues, which sustains a substantial fraction of the HSC pool [47 48 49 50 51 52 ]. Moreover, it has been shown that also neural stem cells localize to vascular niches [53 , 54 ] and that endothelial cells can support the self-renewal of neural stem cells in culture [55 ]. This raises the possibility that endothelial cells are generally important in the formation of mammalian stem cell niches and that sinusoidal endothelium represents a special environment adapted for the maintenance of HSC. Our findings that Sca-1 is expressed in CD45+ skin cells and CD45–CD31+ vascular endothelial cells further support such a relationship (data not shown). Thus, it is possible that the Sca-1+ skin cell subpopulations contribute to maintaining precursor pools in the skin and might even constitute together the stem cell niches within the differentiated tissue.

As we identified only minute numbers of donor-derived cells in the skin, it is conceivable that upon arrival in their niche, the stem cells remain in a relatively quiescent state as previously reported for HSC in the BM [22 , 56 ]. It would be interesting to identify the signals guiding migration and adhesive interactions of dermal stem/progenitors in the niche of the skin. Our results are similar with the observation that hair follicle dermal cells have hematopoietic activity [4 ], and it remains to be evaluated whether the stem cells/precursors that we have isolated from the whole dermis correspond to hair follicle dermal cells. What remains unclear is the role of HSC in the dermis. It is possible that they provide a means for replenishing tissue cells, whether by fusion with endogenous cells or by direct differentiation into tissue cells. Current concepts suggest that HSC fusion with aging tissue cells is a more likely mechanism [57 ]. To date, there is no evidence that such a process occurs in the dermis, and for the moment, the most apparent role for dermal HSC could be to replenish tissue resident macrophages and dendritic cells.

This study provides the first attempt to phenotype, isolate, purify, and differentiate cells expressing stem cell antigens in the dermis and will facilitate a better characterization of the dermal stem cell molecular signature. Moreover, identification of the precise location of dermal stem cells will ease analysis of the gene-expression profile of the surrounding niche cells. As the stem cell and precursor cell numbers, which can be isolated from the dermis, are extremely low, expansion strategies with a large-scale and long lifespan are needed for application. Nonetheless, results from this report present a platform for further research with great therapeutic potential.


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ACKNOWLEDGEMENTS
 
This project was funded by an Austrian Science Foundation grant (P17078-B14 to A. E-B.). U. S. is supported by a grant from the Deutsche Forschungsgemeinschaft (Schm 2128/1-1). The authors thank Dr. Antal Rot (Novartis Research Institutes for Biomedical Research, Vienna, Austria) for his kind help in preparing the micrographs, Edith Pursch (Novartis Research Institutes for Biomedical Research), Rosaida Alonso-Vasquez and Martina Hammer for technical contributions.

Received January 10, 2006; revised April 20, 2006; accepted May 17, 2006.


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