TISSUE ENGINEERING: Part A Volume 00, Number 00, 2014 ª Mary Ann Liebert, Inc. DOI: 10.1089/ten.tea.2013.0759
Full-Thickness Skin with Mature Hair Follicles Generated from Tissue Culture Expanded Human Cells Xunwei Wu, MD, PhD, Larry Scott Jr., Ken Washenik, MD, PhD, and Kurt Stenn, MD
The goal of regenerative medicine is to reconstruct fully functional organs from tissue culture expanded human cells. In this study, we report a method for human reconstructed skin (hRSK) when starting with human cells. We implanted tissue culture expanded human epidermal and dermal cells into an excision wound on the back of immunodeficient mice. Pigmented skin covered the wound 4 weeks after implantation. Hair shafts were visible at 12 weeks and prominent at 14 weeks. Histologically, the hRSK comprises an intact epidermis and dermis with mature hair follicles, sebaceous glands and most notably, and unique to this system, subcutis. Morphogenesis, differentiation, and maturation of the hRSK mirror the human fetal process. Human antigen markers demonstrate that the constituent cells are of human origin for at least 6 months. The degree of new skin formation is most complete when using tissue culture expanded cells from fetal skin, but it also occurs with expanded newborn and adult cells; however, no appendages formed when we grafted both adult dermal and epidermal cells. The hRSK system promises to be valuable as a laboratory model for studying biological, pathological, and pharmaceutical problems of human skin.
Over the course of a decade, we have tried many systems in an effort to produce full-thickness skin with complete appendages starting with expanded human cells. Recently, we have been able to accomplish this goal. We are now able to reproducibly generate full thickness skin with mature epidermal and dermal layers containing mature shaft producing hair follicles associated with sebaceous glands. Most significantly these constructs produce a full subcutis, and in some cases, eccrine gland structures; we believe these features have not been found in any other previously described skin equivalent. We refer to this construct as human reconstructed skin (hRSK). This report describes the method and morphological characterization of hRSK. hRSK promises to be a valuable tool for future developmental, pathogenetic, and therapeutic studies of mature human skin.
he overarching goal of regenerative medicine is to reform complete functional human organs. Since it is readily accessible and relatively simple in structure and function, skin has been an early target in the field. In fact, the first clinical successes in tissue-engineered products have been skin substitutes made of collagen sheets with embedded fibroblasts alone1 or with overlaid epidermal cells.2 Surgeons have used these equivalents to cover cutaneous wounds due to trauma, burn, surgery, or pathogenic ulceration. Nevertheless, they have had limited long-term clinical application because they lack normal dermal structure, subcutis, and appendages. Heretofore, workers in the field have not been able to form a mature skin construct with skin appendages when starting with tissue culture expanded human cells.3–6 Greater success has been realized using fresh mouse cells alone or in combination with human cells.5,7–10 Recently, Li et al.11 and Thangapazham et al.12 using tissue culture expanded TSC2 null fibroblasts or normal dermal papilla cells plus human newborn foreskin keratinocytes were able to reconstruct a human skin equivalent in vivo. Whereas this skin showed normal epidermal differentiation and hair follicle formation, it did not form eccrine structures, arrector pili muscles, or outgrowing hair shafts.11,12 The same lack of complete hair follicle maturation was reported by Higgins et al., who used dermal papilla cell spheres combined with newborn foreskin epidermal sheets.13
Methods Cell isolation and culture
Epidermal and dermal cells were isolated from human fetal scalp tissues [estimated gestational age (EGA) 15–18 weeks. ABR (Advanced Bioscience Resources, Inc.); the Institutional Review Board (IRB) review provided by ABR] following the procedure of Zheng et al.14 Briefly, the skin was cut into 5 · 5 mm strips, and incubated in phosphatebuffered saline (PBS) Dispase (2.5 mg/mL; Sigma Chem. Co) overnight at 4C. The next day, the epidermis and dermis were separated using forceps. For dermal cell preparations, the
Aderans Research Institute, Inc., Marietta, Georgia.
dermis was minced using crossed scalpel blades and incubated in collagenase (2.5 mg/mL, 37C). After 30 min, the dermal solution was neutralized in Dulbecco’s modified Eagle’s medium (DMEM) with 10% fetal bovine serum (FBS), passed through 100 mm filter, and the filtrate centrifuged and rinsed with PBS. The pellet of dermal cells was plated in DMEM/F12 (3:1) containing 0.1% penicillin/streptomycin, 40 mg/mL fungizone, 40 ng/mL FGF2, 20 ng/mL EGF, 2% B27 supplement with 5% FBS; medium was changed every 5 days. Dermal cells were passaged when the cell density attained 80–90% confluence; passage dilution was 1:5–10. For epidermal cell preparations, cells were collected from the isolated epidermis after trypsin incubation (porcine; Sigma Chem. Co., Inc. 0.025% in PBS, 37C, with shaking, 30 min), neutralized with 10% FBS in DMEM, filtered (100 mm filter; Millipore), centrifuged, and PBS rinsed. The epidermal cells were plated into culture vessels containing CNT07 (CELLnTEC) plus Rock inhibitor Y27632 (Sigma Chem. Co; 10 mM/mL). After 3 days culture, Y27632 was removed; medium was changed every other day. Epidermal cells were passaged when the cell density attained 80–90% confluence, and the passage dilution was 1:3. Both dermal and epidermal cells were collected for grafting after the density reached 90–100% confluence. To allow adequate expansion, all cells used for grafting were passaged from one to four times. Epidermal cells were isolated from neonatal human foreskins, (purchased from Cooperative Human Tissue Networks, University of Alabama, Birmingham) similar to the method used for fetal epidermal cells. Since the efficiency of hRSK formation by newborn foreskin keratinocytes was similar to that generated by fetal epidermal cells and because it was easy to obtain, we used foreskin keratinocytes as the main epidermal cell source for all our experiments. Adult male dermal and epidermal cells were isolated from full-thickness scalp skin (human donor, Quorum Review IRB, Seattle, WA, approved protocol). Briefly, adult scalp tissue was chopped by scalpel blades, and incubated in the digestion solution (2.5 mg/mL dispase and 2.5 mg/mL collagenase in DMEM) at 37C; after 1 h, trypsin (0.05%) was added to the solution for another half hour. The digested tissue preparation was passed through a 100 mm filter and then through a 40 mm filter. The filtrate was used for epidermal cell preparations and the retentate for dermal cell preparations. The adult dermal and epidermal cells were cultured separately. The culture conditions for adult and fetal cells were the same. Grafting dissociated human cells onto skin wounds of immunodeficient mice
For grafting of the tissue culture expanded cells, we used a modified method described by Lee et al.10 Briefly, 2–4 · 106 dissociated human epidermal cells were mixed with 3–6 · 106 human dermal cells (the ratio of epidermal and dermal cells was range 1:1.5–2) to make a cell slurry in a total volume of 150 mL of DMEM/F12 (1:1) (Life Technologies; # 11039). The cell slurry was transferred onto a PET membrane (polyethylene terephthalate, 3.0 mm pore size), which was excised from a six-well cell culture insert (Becton Dickinson; Cat. No. 353091), and incubated at 37C for 1–1.5 h in a tissue culture incubator. The membrane with non-adherent cell slurry was flipped upside down onto a back skin 1–2 cm2 wound created either on a nu/nu or NOD/SCID mouse (Charles River Lab) (Although we used two different immunodeficient mice, we
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found in earlier studies that both strains worked equally well). The size of the excision wound was determined by the size of the cell slurry. The membrane was sutured to the host skin, covered with Vaseline (Unilever; Englewood) and then wrapped by sterile tape (Supplementary Fig. S1A; Supplementary Data are available online at www.liebertpub.com/tea). The dressings were removed after 1 week and the PET membrane shed as the outer epidermis formed. Each mouse bore either one or two grafts. Mouse work was performed in accordance with protocols approved by the Institutional Animal Care and Use Committee (IACUC) of Saint Joseph’s Translational Research Institute, Atlanta, GA. Histology and immunofluorescence analysis
Tissues were embedded in Optimal Cutting Temperature Compound (OCT; Tissue-Tek, Sakura Finetek) and frozen; 10 mm cryosections were taken for histological analysis following standard protocols.15 Immunohistochemical analysis was performed as following: the cryosection was fixed in 4% paraformaldehyde, and incubated with the blocking buffer (2% bovine serum albumin and 5% donkey serum in PBS with 0.01% Triton · 100) at room temperature for 1 h, then applied with primary antibody at 4C overnight. On the second day, after removing the primary antibody solution, the specimen was incubated with secondary antibody at room temperature. After 1 h incubation, the section was washed with PBS and mounted (Mounting Medium with DAPI; Vector, H-1200), and the staining was analyzed by confocal microscope. The following primary and secondary antibodies and dilution times (antibodies were diluted in blocking buffer) were used: rat anti-FITC conjugated (CD49) a6-integrin (Stem Cell; Cat. No. 10111, 1:100), mouse anti-human pan-cytokeratin (BD; Cat. No. 550951, 1:400); polyclonal rabbit anti-keratin 5 (Covance; Cat. No. PRB-160p, 1:400); polyclonal rabbit antikeratin 10 (Covance; Cat. No. PRB-159p, 1:400); monoclonal mouse anti-keratin 17 (Serotec; Cat. No. MCA1872, 1:200); monoclonal mouse anti-keratin 19 (Santa Cruz; Cat. No. SC-6278, 1:400); polyclonal rabbit anti-keratin 6 (Abcam; Cat. No. Ab75703, 1:200); monoclonal mouse anti-vimentin (5G3F10)(Cell Signaling; Cat. No. 3390, 1:200); mouse antihuman nuclei antibody (Millipore; Cat. No. MAB1281, 1:200); rabbit monoclonal antibody anti-mouse CHD3 (Epitomics; Cat. No. 3294-1, 1:200); The following secondary antibodies (all from Life Technologies) were used: Alexa fluor-488 donkey anti-mouse IgG (Cat. No. A21202); Alexa fluor-594 donkey anti-mouse IgG (Cat. No. A21203); Alexa fluor-488 donkey anti-rabbit IgG (Cat. No. A21206); Alexa fluor-594 donkey anti-rabbit IgG (Cat. No. A21207). 1:400 dilution was used for all secondary antibodies. We demonstrated the human versus mouse specificity by testing all these antibodies on both mouse and human tissues. Alkaline phosphatase staining
Alkaline phosphatase (AP) staining was carried out according to the manufacturer’s protocol (Roche (NBT/BCIP stock solution; Cat. No. 11 681 451 001). Briefly, cryosections were rinsed in PBS, fixed in ice cold acetone for 10 min, then washed 2 · 5 min in 1· PBS solution. The sections were placed in the AP buffer (0.1 M Tris-HCL, PH 9.5, 0.1 M NaCl, 0.05 M MgCl2) for 5 min and then incubated in freshly prepared staining solution (200 mL NBT/BCIP stock solution
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in 10 mL AP butter) for 5–30 min at room temperature. After dark blue staining appeared, the specimens were washed, counterstained (eosin for 30 s), dehydrated with alcohol, and coverslip mounted. Results Full-thickness hair-producing skin from tissue culture expanded human epidermal and dermal cells–gross observations
In these studies, the dermal cells were prepared from eight different fetal scalp specimens and the epidermal cells from six different foreskins. Four weeks after implanting, the tissue culture expanded cells formed a pigmented skin closure to the wound surface (Supplementary Fig. S1B). The hRSK covered an area ranging from 0.1 to 2 cm2 (Supplementary Fig. S1B and Fig. 1A, B). Hair shafts visible at 12 weeks were prominent at 14 weeks (Fig. 1A-A¢ and Supplementary Fig. S1B). Of the 18 grafts implanted, 15 (83%) formed hair follicles and the number of shafts per hRSK ranged from 20 to 1000 per cm2 (Fig. 1A-A¢, B, C, and Supplementary Fig. S1B). There was no appreciable difference in resultant hRSKs when we used cells derived from fetal tissues of different EGA. Six months after grafting, resultant shafts were 2–3 cm in length (Fig. 1C). Although we followed hair growth in this model for up to 1 year, we did not find hair shafts longer than 3 cm. We estimate that the anagen growth phase in the hRSK lasts 2–3 months. We ascribe the variation, in response between preparations, to the heterogeneity of the human donor population sampled. To further assess the regenerative ability of hRSK follicles, we plucked the outgrowing hair shafts from three constructs. After this stimulus, hair regrowth resumed producing shafts at a rate of about 1 cm of linear growth per month (Supplementary Fig. S1C).
3 Implanted cells undergo hRSK skin regeneration reminiscent of normal human embryonic morphogenesis–histological observations
Taking a histological approach, we differentiated the stages of hRSK morphogenesis. At 1 week after cell implantation, a thin skin, containing two layers of epidermis and dermis, was already formed (Fig. 2A), but at this time point, no mesenchymal cells express AP (Fig. 2B). At 2 weeks, the basal epidermal cells became more organized and polarized. At 3 weeks, epidermal placodes formed (Stage 1 follicle). Shortly after, dermal cell condensates collected below the placodes (Fig. 2A), which expressed AP (Fig. 2B). The sequence of events observed in the hRSK recapitulated our earlier observations, which defined the pathway dissociated cells undergo to form skin.14 Mature hRSK consists of epidermal, dermal, and subcutis layers with appendages: hair follicles, sebaceous, and eccrine glands–histological observations
At 10 weeks the histology of hRSK mirrors the structure of 18 weeks old (EGA) human fetal scalp skin (Supplementary Fig. S2A). At 6 months the interfollicular epidermis of the hRSK (Supplementary Fig. S2A and Fig. 3A) has a defined rete pattern and fully formed basal, spinous, granular, and corneal layers (Supplementary Fig S2B). The basal layer of the reformed epidermis is well-pigmented suggesting normal melanogenesis (Supplementary Fig. S2B). The hRSK dermis at 6 months has a compact superficial layer and a loosely structured deep layer. Most surprising to us was that at this time the hRSK contained a prominent subcutis made of mature adipose tissue (Fig 3A and Supplementary Fig. S2A, C); subcutis appeared in all hRSKs, showing hair follicle regeneration. Immunohistochemistrical analysis demonstrated that the hRSK epidermis expressed
FIG. 1. Skin regeneration (hRSK) from dissociated and tissue culture expanded human epidermal and dermal cells. (A) hRSK with numerous hair shafts at 3 months after grafting dissociated human cells. (A¢) is a higher magnification image of (A, B) hRSK covers about 2 cm2 (2 months after cell grafting). (C) Regenerated hair shafts, *3 cm long (6 months after grafting). Scale bar: 0.5 cm.
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FIG. 2. Formation of hair follicles in the hRSK recapitulates fetal morphogenesis. hRSK were harvested at 1, 2, 3, and 4 weeks after cell grafting and stages of morphogenesis assessed. (A) Placode formation was first found at 3 weeks (black arrows), with stage 2–3 follicles forming at 4 weeks (H & E histology); (B) by alkaline phosphatase (AP) staining the first signs of dermal condensation occurred at 3 weeks (black arrows), with strong dermal AP expression at 4 weeks. Scale bar: 50 mm.
expected differentiation markers described by previous workers,16 namely, basal keratinocytes express keratins, K5 and K14, and the suprabasal cells express differentiation-related keratins, K1 and K10 (Supplementary Fig. S3A, B and data not shown). The a6 integrin staining indicated an intact basement membrane formed at the junction of the epidermis and dermis in hRSK (Supplementary Fig. S3A–D). The follicles were for the most part terminal in size, although they were smaller compared with normal scalp follicles. Although most follicles were anagen in growth phase, some constructs contained catagen and telogen forms (Fig. 3A and Supplementary Fig. S4A). Anagen follicles showed characteristic outer root sheath and inner root sheath layers (Fig. 3B, C). By immunohistochemistry the hair follicle sheaths, but not the epidermis, expressed K17 (Supplementary Fig. S3C) and K6 (Supplementary Fig. S3D), which are follicular keratins.16 All regenerated follicles were associated with mature sebaceous glands (Fig. 3A, B, and Supplementary Fig. S4B) and arrector pili muscles (Fig 3B, D, E). Each defined bulb in the proximal anagen follicle clasped an intact dermal papilla (Fig. 3A, C, F, G). The hair shafts were pigmented and had typical human shaft features with thick solid cortex and absent to rare medulla; ladder shaped medullae typical of the mouse were not seen in the hRSKs (Supplementary Fig. S4A). Follicle growth phases as assessed by trichogram, a conventional shaft plucking method of analysis, confirmed anagen, catagen, and telogen shaft forms (Fig. 3H). Quantification data reveal that 15% of plucked hairs showed telogen morphology, and the remaining were anagen (5 months), a ratio typical of a normal human scalp trichogram17 (Fig. 3I). These observations suggest that there is a population of cycling hair follicles in the hRSK. In some cases (around 5% of hRSK), eccrine structures were present in the deep dermis and subcutis with glandular, straight and coiled ductal portions; the ducts appeared to extend from the subcutis to the epidermis (Fig. 3J and
Supplementary Fig. S4C). The eccrine ducts showed characteristic K19 positive staining (Fig. 3K). Collectively, these results indicate that hRSK contains normally differentiating epidermis, dermis, subcutis, pilosebaceous apparatus and in some cases, eccrine structures. The regenerated skin and its appendages are of human origin–histological study
To confirm that the structures we observed arose from the implanted human cells, we used human specific antibodies. To check the claimed specificity of the antibodies, we carefully stained both mouse and human tissues with each antibody and found no cross-species staining (e.g., white arrows Fig. 4A; white arrows left panel Fig. 4E, G and Supplementary Fig. S5). At 6 weeks and 6 months hRSK expressed the human specific antigen pan-cytokeratin (Fig. 4A) in the interfollicular epidermis and early stage follicles (Fig. 4B). To test the dermal cells, we used human specific vimentin antibody, a mesenchymal cell marker.19 At 6 weeks and 6 months dermal stromal cells (Fig. 4C, D) and the dermal papilla (arrowhead, Fig. 4D, G), dermal sheath (Fig. 4D, G) and the subcutis (Supplementary Fig. S5G) expressed these markers. Double staining with pan-cytokeratin (red) and vimentin (green) confirmed the conclusion that the cells making up the hRSK are human (Fig. 4G). Finally, anti-human nuclei antibody, a marker, which recognizes all human cellular nuclei, stained both epidermal and dermal tissues of the hRSK, but not the mouse tissues (Fig. 4E, F, and Supplementary Fig. S5A–F). Next we probed the extent of mouse cell infiltration into the hRSK. Using an antibody specific to mouse CHD3, a member of the chromodomain helices DNA-binding (CHD) family of proteins, we found very few CHD3 positive mouse cells in the dermis or epidermis of 6 weeks hRSK; staining controls showed that mouse tissues stained, whereas
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FIG. 3. The hRSK showing full-thickness skin with mature hair follicles, sebaceous glands, dermal papillae, and eccrine glands. (A) Histology of 6 months hRSK showing full-thickness skin. Scale bar: 500 mm. (B, C) Regenerated follicles with outer root sheath (ORS), inner root sheath (IRS), arrector pili muscle (APM), sebaceous gland (SB), dermal sheath (DS), and dermal papilla (DP). Scale bar: 100 mm. (D) Arrector pili muscle (black arrows). (E) a-smooth muscle actin (a-SMA) antibody (green) confirms APM. (F–G) Alkaline phosphatase (AP, blue) stains dermal condensate at 6 and 10 weeks (H) Plucked hair shafts from 5 months hRSK. (I) Percent of telogen shaft forms increase from 3 to 7 months (per 60–90 total number of plucked shafts, n = 3); ( J) Eccrine structure, gland, and duct in the deep dermis in 3 months hRSK. (K) K19 (red) staining of eccrine ducts; basement membrane defined by a6 integrin (a6, green). (F–K). Scale bar: 50 mm.
human tissues did not (Supplementary Fig. S5D–F). At 6 months (Fig. 4H, right panel), we found a few CHD3 positive (red) cells in the dermis, but not in the epidermis. Quantification of CHD3 positive cells in the dermis of hRSK showed that the percentage of mouse cells in the construct at 6 weeks and 6 months was around 2% and 9% of all DAPI positive cells, respectively. This study suggests that there is a predominance and persistence of human cells in the hRSK, but there is also mouse cell infiltration which increases with time. Although we observed human cell antigen retention for up to 6 months, further studies will be needed to establish how long human cells persist in this model.
Cultured adult cells when combined with younger cells were able to regenerate hRSK with hair follicles, but less efficiently
In all of the constructs reported above, the skin cells arose from either fetal or newborn human tissues. Next, we asked whether cultured adult cells could regenerate skin with hair follilces like the cells from younger individuals. We found that adult cells were able to generate hair follicles associated with sebaceous glands when combined with either fetal or newborn cells, but the efficiency of forming hair follicles was less compared with fetal cells (Fig. 5A–C, A¢–C¢). When adult dermal cells were combined with adult epidermal cells,
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FIG. 4. The hRSK is predominately of human cell origin. (A, B) Human pan-cytokeratin (pan-ck) present in the epidermis and ORS; white arrow margin of human and mouse skin. (C, D) Human vimentin in dermis at 6 weeks (red) and 6 months (green); dermal papilla (arrowhead). (E, F) Human nuclei antigen stain; mouse cells do not stain (white arrows). (G) Double staining with pan-cytokeratin (red) and vimentin (green) at 6 and 12 weeks; white arrow indicates both antibodies did not stain mouse tissue. Arrow head DP at 12 weeks. (H) Immunostaining for mouse specific antibody CHD3 (red), DAPI for nuclei (blue); more CHD3 positive cells infiltrate the hRSK dermis at 6 months. The epidermal– dermal junction by dashed lines (D) red, (F) green, (H) green), a6 integrin (A–C, E) green), DAPI for nuclei (blue). (A– H). Scale bar: 50 mm. although a skin-like cover formed, appendages did not (0%, 0 out of 9) (Fig. 5D, D¢). These data indicate cell donor age is important and that appendage formation requires a property which adult cells lose. Discussion
Starting with tissue culture expanded cells isolated from human epidermal and dermal tissues and placed within a mouse skin wound, we have been able to generate fullthickness, appendage-rich skin, which we refer to as hRSK. In its formation, the hRSK undergoes morphogenetic stages reminiscent of developing human fetal skin. The fully formed
construct shows epidermis, dermis and, most unique to this system, a subcutis. The follicles that formed show normal sheath structure with sebaceous glands, muscle, and fully formed outgrowing pigmented shafts. These properties fulfill, in part, the properties of a reengineered follicle as defined by Chuong et al.18 Cell surface marker studies confirm that the cells making up the hRSK are human and that these markers remain stable for at least 6 months. Based on many years of study and compared with the other assays, we believe the success of our effort to produce the hRSK is possibly due to at least two factors. First, cell culture conditions. The culture methods we used appear to have stabilized the skin/hair follicle potential of the cells.
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FIG. 5. Donor age affects appendage formation in the hRSK. Representative images of hRSK (A–D) and its correlated histology (HE staining, A¢–D¢) from different cell combinations. (A, A¢) hRSK with hairs formed from foreskin epidermal cells (FK Epi) with fetal dermal (fDer) cells showed that 15 out of 18 constructs produced hair. (B, B¢) hRSK with hairs formed from foreskin epidermal cells (FK Epi) with adult dermal (Adult Der) cells showed that 5 out of 15 constructs produced hair. (C, C¢) hRSK with hairs formed from adult epidermal cells (Adult Epi) with fetal dermal (fDer) cells showed that 2 out of 17 constructs produced hair. (D, D¢) hRSK with no hairs formed from adult epidermal (Adult Epi) with adult dermal (Adult Der) cells showed that 0 out of 9 constructs produced hair. All data were taken from 3 months old hRSK. Scale bar: (A–D) 250 mm; (A¢–D¢) 100 mm.
For instance, we grew epidermal cells in the presence of the ROCK inhibitor, Y-27632, which reportedly preserves stem cell properties of epidermal cells.20,21 We believe that the use of ROCK inhibitor in our culture plays an important role for the regeneration of hRSK, but the underlying mechanism needs to be studied further. The cells we used for both the epidermal and dermal preparations came from batch preparations where cell population selection was a result of tissue culture conditions and not cloning. Clearly, the cells selected for have skin stem cell and trichogenic potential; in fact when tested, our isolated dermal cell population showed prototypical properties of mesenchymal stem cells (Marshall, Wu, Watkins, and Stenn, unpublished). Cell cloning and better characterization of the important primary cell populations are critical goals of future work. Second, cell placement. We believe the experiments are telling us that how cells are placed is very important to the process of normal organogenesis. The apparently unstructured slurry may allow for mesenchymal-epidermal interactions, which enhance tissue self-organization. From the frustrated attempts of other workers3,5,22 as well as our own, we have concluded that if the cells are not restricted, but allowed to organize themselves, without imposing a confining scaffold, a more complete regeneration will result. We realize this conclusion contradicts traditional dogma23,24 which prescribes cell delivery with an appropriate matrix, or scaffold. Our own work, as well as the work cited by Sasai25 and recent report from Takebe,26 suggests that the proper cells will produce their own support structure as they undergo the morphogenetic process.23 We would assert that the emphasis in regenerative work should be on the cells and less on
the scaffold/matrix, which the investigator might believe the cells need to do their job. In this study, we used fetal and newborn human foreskin epidermal cells and fetal and adult scalp dermal cells. When the epidermal and dermal cells were both derived from fetal skin, hRSK reformation was optimal. When the epidermal component arose from fetal or newborn skin and the dermal from adult skin, the hRSK formed was less appendage rich. When both the epidermal and dermal cells arose from the adult skin, no appendages formed. This result underscores the importance of donor age and puts the focus on tissue aging as an important parameter. Clearly, as we try to bring these constructs to the clinic, we will have to elucidate the basis for this difference. Although the mouse as a model for human skin and its diseases has been powerfully instructive, there comes a point when such studies must be done on human tissues. Having a mature full-thickness human skin model for study will be relevant to the study of uniquely human problems. This system will be valuable for example, in morphogenetic lineage studies of human skin appendages, in skin disease models where cells from diseased skin could be used to generate the hRSK, in testing hypotheses regarding evolution of human skin differences, and in testing therapeutic agents and regimens for efficacy and toxicity well before clinical studies. We would hope that the lessons learned while working with this model will give insights into new approaches for treating chronic skin ulcers and burns. Finally, we would predict that aspects of this model will further the understanding and development of scarfree wound repair.
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No competing financial interests exist. References
1. Naughton, G.K., and Mansbridge, J.N. Human-based tissueengineered implants for plastic and reconstructive surgery. Clin Plast Surg 26, 579, 1999. 2. Parenteau, N. Skin: the first tissue-engineered products. Sci Am 280, 83, 1999 3. Michel, M., et al. Characterization of a new tissue-engineered human skin equivalent with hair. In Vitro Cell Dev Biol Anim 35, 318, 1999. 4. Havlickova, B., Biro, T., Mescalchin, A., Arenberger, P., and Paus, R. Towards optimization of an organotypic assay system that imitates human hair follicle-like epithelialmesenchymal interactions. Br J Dermatol 151, 753, 2004. 5. Larouche, D., Cuffley, K., Paquet, C., and Germain, L. Tissue-engineered skin preserving the potential of epithelial cells to differentiate into hair after grafting. Tissue Eng Part A 17, 819, 2011. 6. Krugluger, W., et al. Reorganization of hair follicles in human skin organ culture induced by cultured human folliclederived cells. Exp Dermatol 14, 580, 2005. 7. Toyoshima, K.E., et al. Fully functional hair follicle regeneration through the rearrangement of stem cells and their niches. Nat Commun 3, 784, 2012. 8. Sriwiriyanont, P., et al. Morphogenesis of chimeric hair follicles in engineered skin substitutes with human keratinocytes and murine dermal papilla cells. Exp Dermatol 21, 783, 2012. 9. Ehama, R., et al. Hair follicle regeneration using grafted rodent and human cells. J Invest Dermatol 127, 2106, 2007. 10. Lee, L.F., Jiang, T.X., Garner, W., and Chuong, C.M. A simplified procedure to reconstitute hair-producing skin. Tissue Eng Part C Methods 17, 391, 2011. 11. Li, S., et al. Human TSC2-null fibroblast-like cells induce hair follicle neogenesis and hamartoma morphogenesis. Nat Commun 2, 235, 2011. 12. Thangapazham, R.L., et al. Dissociated human dermal papilla cells induce hair follicle neogenesis in grafted dermalepidermal composites. J Invest Dermatol 134, 538, 2014. 13. Higgins, C.A., Chen, J.C., Cerise, J.E., Jahoda, C.A., and Christiano, A.M. Microenvironmental reprogramming by three-dimensional culture enables dermal papilla cells to induce de novo human hair-follicle growth. Proc Natl Acad Sci U S A 110, 19679, 2013. 14. Zheng, Y., et al. Mature hair follicles generated from dissociated cells: a universal mechanism of folliculoneogenesis. Dev Dyn 239, 2619, 2010. 15. Presnell, J.K., Schreibman, M.P, and Humason, G.L. Humason’s Animal Tissue Techniques. Baltmore, MD: John Hopkins University Press, 1997, p. 572.
16. Moll, R., Divo, M., and Langbein, L. The human keratins: biology and pathology. Histochem Cell Biol 129, 705, 2008. 17. Dawber, R.P. The embryology and development of human scalp hair. Clin Dermatol 6, 1, 1988. 18. Chuong, C.M., Cotsarelis, G., and Stenn, K. Defining hair follicles in the age of stem cell bioengineering. J Invest Dermatol 127, 2098, 2007. 19. Goodpaster, T., et al. An immunohistochemical method for identifying fibroblasts in formalin-fixed, paraffin-embedded tissue. J Histochem Cytochem 56, 347, 2008. 20. Chapman, S., Liu, X., Meyers, C., Schlegel, R., and McBride, A.A. Human keratinocytes are efficiently immortalized by a Rho kinase inhibitor. J Clin Invest 120, 2619, 2010. 21. Terunuma, A., Limgala, R.P., Park, C.J., Choudhary, I., and Vogel, J.C. Efficient procurement of epithelial stem cells from human tissue specimens using a Rho-associated protein kinase inhibitor Y-27632. Tissue Eng Part A 16, 1363, 2010. 22. Limat, A., Hunziker, T., Breitkreutz, D., et al. Organotypic cocultures as models to study cell-cell and cell-matrix interactions of human hair follicle cells. Skin Pharmacol 7, 47, 1994. 23. Wood, F. Tissue engineering of skin, chapter 57. In: Atala, A., Lanza, R., Thompson, J.A., and Nerem, R., eds. Principles of Regnerative Medicine, 2nd edition. Elsevier, Inc., 2011. 24. Mahjour, S.B., Ghaffarpasand, F., and Wang, H. Hair follicle regeneration in skin grafts: current concepts and future perspectives. Tissue Eng Part B Rev 18, 15, 2012. 25. Sasai, Y. Cytosystems dynamics in self-organization of tissue architecture. Nature 493, 318, 2013. 26. Takebe, T., Sekine, K., Enomura, M., Koike, H., Kimura, M., Ogaeri, T., Zhang, R.R., Ueno, Y., Zheng, Y.W., Koike, N., Aoyama, S., Adachi, Y., and Taniguchi, H. Vascularized and functional human liver from an iPSC-derived organ bud transplant. Nature 499, 481, 2013.
Address correspondence to: Kurt Stenn, MD 46 Bayard Lane Princeton, NJ 08540 E-mail: [email protected]
Xunwei Wu, MD, PhD 30 Revere Beach Parkway Unit 607 Medford, MA 02155 E-mail: [email protected]
Received: December 18, 2013 Accepted: June 13, 2014 Online Publication Date: August 5, 2014