Chapter 3.4
Epidermal structure created by canine hair follicle keratinocytes enriched with bulge cells in a three-dimensional skin equivalent model in vitro: implications for regenerative therapy of canine epidermis
Background – Keratinocytes in the hair follicle bulge region have a high proliferative capacity, with characteristics of epithelial stem cells. This cell population might thus be an ideal source for generating the interfollicular epidermis in a canine skin equivalent.
Hypothesis/Objectives – This study was designed to determine the ability of canine hair follicle bulge cell-enriched keratinocytes to construct canine living skin equivalents with interfollicular epidermis in vitro.
Animals – Four healthy beagle dogs from a research colony.
Methods – Bulge cell-enriched keratinocytes showing keratin 15 immunoreactivity were isolated from canine hair follicles and cultured on dermal equivalent containing canine fibroblasts. Skin equivalents were subjected to histological, immunohistochemical, western blot and RT-PCR analyses after 10-14 days of culture at the air-liquid interface.
Results – The keratinocyte sheets showed an interfollicular epidermal structure comprising four to five living cell layers covered with a horny layer. Immunoreactivities for keratin 14 and desmoglein 3 were detected in the basal and immediate suprabasilar layers of the epidermis, while keratin 10 and desmoglein 1 occurred in more superficial layers. Claudin 1 immunoreactivity was seen in the suprabasalar layer of the constructed epidermis, and filaggrin monomers and loricrin were detected in the uppermost layer. Basal keratinocytes in the skin equivalent demonstrated immunoreactivity to antibodies against basement membrane zone molecules.
Conclusions and clinical importance – A bulge stem cell-enriched population from canine hair follicles formed interfollicular epidermis within 2 weeks in vitro, and thus represents a promising model for regenerative therapy of canine skin.
Introduction
The skin is the outermost surface of the animal’s body and plays a crucial role in providing a mechanical and biological barrier against microbial organisms, chemical components and ultraviolet light. The skin also protects the body from dehydration and maintains the physiological environment inside the body. Large skin defects caused by extensive wounding, trauma, burns, chronic ulcers or surgical procedures involved in removing large tumours are major concerns in veterinary medicine.
Wound healing is a complex process that includes inflammation, angiogenesis, re-epithelialization and remodeling of the extracellular matrix processes.1,2 Re-epithelialization is a crucial step in wound healing, and the development of a new therapeutic approach to accelerate re-epithelialization is a key goal for successful wound management.3 It is notable that hair-covered areas recover sooner than hairless areas following skin injury, as re-epithelialization spreads from the hair follicle infundibulum.4,5 This suggests that hair follicle keratinocytes may contribute to re-epithelialization of the interfollicular epidermis.
The hair follicle bulge region constitutes the outermost layer of the outer root sheath and the attachment site of the arrector pili muscle. In addition, it was recently reported to harbour epithelial stem cells in mice.6–8 This cell population has been shown to contribute to the regeneration of the interfollicular epidermis after wounding.9 These findings suggest that hair follicle stem cells may represent a promising target for the development of new treatment approaches for skin wounds. The bulge region in canine hair follicles contains keratin 15 (K15)-positive keratinocytes with characteristics of epithelial stem cells.10,11 These K15-positive cells have a greater proliferative capacity than interfollicular epidermal keratinocytes in vitro, and a better ability to reconstitute pilosebaceous units and interfollicular epidermis in vivo.11 Canine hair follicle bulge cells may therefore represent a more effective cell source for the regeneration of skin wounds than canine interfollicular epidermal keratinocytes.
Three-dimensional (3D) skin equivalents have been developed as ex vivo dermatological research materials12 and as new therapeutic materials for wound treatment.13,14 Keratinocytes in these 3D skin equivalents can recapitulate in vivo epithelial cell differentiation in vitro and generate stratum corneum and the dermoepidermal junction.12 These 3D skin equivalents are considered to possess similar physiological properties to normal skin. The goal of the present study was to investigate the potential of canine hair follicle bulge cells to construct interfollicular epidermis in skin equivalents and assess the ability of the cell population to differentiate into intact interfollicular epidermis in vitro.
Materials and methods
Skin samples
Three healthy beagle dogs (3-4 years old) were used in this study. Keratinocytes and fibroblasts were isolated from skin samples obtained from the dorsolateral skin (20 mm × 20 mm) after sedation with medetomidine hydrochloride (20-80 μg/kg; Domitor; Pfizer Japan, Tokyo, Japan) and local anaesthesia with lidocaine hydrochloride (Xylocaine; AstraZeneca Japan, Osaka, Japan). For immunohistochemical staining of normal canine skin, samples were obtained from the nose and footpads of a healthy beagle dog using a skin biopsy punch after euthansia for reasons unrelated to this study, and were embedded in optimal cutting temperature (OCT) compound (Sakura Finetek, Tokyo, Japan) or fixed with 10% neutral buffered formalin for paraffin embedding. All experimental procedures were officially approved by the Animal Research Committee and were carried out in accordance with the ethical guidelines of Tokyo University of Agriculture and Technology.
Isolation and culture of bulge cell-enriched keratinocytes
Canine bulge cell-enriched keratinocytes, which were positive for the bulge stem-cell marker K15, were isolated from hair follicles as previously described.10,11 Skin samples were cut vertically into small pieces. The middle parts of the fragments, containing the isthmus and suprabulbar parts of the hair follicles, including the bulge region,10 were carefully microdissected and incubated with 1500 U/mL of dispase II (Godo Syusei, Tokyo, Japan) overnight at 4°C. Hair follicle epithelia were separated manually from the adjacent dermis and incubated with 0.05% trypsin EDTA for 15-20 min at 37°C to isolate single hair follicle keratinocytes. Isolated cells were filtered using a 100 μm cell strainer (Becton Dickson, Franklin Lakes, NJ, USA) and counted.
Isolated keratinocytes were cultured in William’s medium (Invitrogen, Carlsbad, CA, USA) supplemented with 10% NuSerum lV (Becton Dickson), 5 ng/mL epidermal growth factor (Sigma-Aldrich, St Louis, MO, USA) and 10−10 mol/L cholera toxin (List Biological Laboratories, Campbell, CA, USA) on type 1 collagen-coated dishes (Iwaki, Chiba, Japan) at 37°C in a humidified atmosphere of air supplemented with 5% CO2 as previously reported.10,11 Culture medium was changed every 3-4 days. Cultured keratinocytes at passages two to three were stored in liquid nitrogen until use.
Isolation and culture of canine dermal fibroblasts
Canine dermal pieces comprising 2 mm cubes were placed on culture dishes and incubated with Dulbecco’s modified Eagle’s medium (DMEM; Invitrogen) containing 10% fetal bovine serum (FBS) at 37°C in a humidified atmosphere of air supplemented with 5% CO2. Outgrowth of fibroblasts from the dermal pieces was observed within 2 weeks. Canine dermal fibroblasts at passages two to four were used in the following experiments.
Establishment of canine skin equivalent using bulge cell-enriched keratinocytes
Canine fibroblasts (1 × 105 cells/mL) were suspended in FBS. Collagen gel solution was made, containing a mixture of bovine collagen type I (Cellmatrix; Nitta Gelatin Inc., Osaka, Japan), 5x-concentrated DMEM (Cellmatrix), 10x-concentrated Ham’s F-12 medium (Cellmatrix) and fibroblast-containing FBS at a ratio of 7:1:1:1. The collagen gel solution was placed on silicon sheets in six-well culture plates, and incubated at 37°C in a humidified atmosphere of air supplemented with 5% CO2 for 1 h until the solution gelled to form a dermal equivalent. The dermal equivalent was then detached from the silicon sheet and cultured for a further 2-3 days.
The canine dermal equivalent was placed on stainless-steel mesh (0.29 mm mesh) to avoid further shrinkage of the gel. A polycarbonate ring was placed on the dermal equivalent, and 4 × 105 keratinocytes were added into the ring. Fresh culture medium [DMEM containing 10% FBS, 10−10 mol/L cholera toxin (List Biological Laboratories), 10 ng/mL epidermal growth factor (Sigma-Aldrich), 0.4 μg/mL hydrocortisone (Sigma-Aldrich), 5 μg/mL transferrin (Sigma-Aldrich), 10−8 mol/L retinoic acid (Sigma-Aldrich), 2 nmol/L 3,3′-5 tri-iodothyronine (Sigma-Aldrich), 5 μg/mL insulin (Sigma-Aldrich), 100 U/mL penicillin, 100 μg/mL streptomycin and 0.25 μg/mL amphotericin (CellnTEC, Bern, Switzerland)] was applied outside the ring. The ring was removed after 1-2 days, and 2 mL of fresh culture medium was added into the dish to start the cell culture at the air-liquid interface. Cells were incubated at 37°C, in a humidified atmosphere of air supplemented with 5% CO2, and the culture medium was changed every 3 days, with the addition of 10 μmol/L matrix metalloproteinase lll inhibitor (Merck Millipore, Billerica, MA, USA) and 1.5 μmol/L aprotinin (Roche Diagnostics, Almere, The Netherlands) 8 days after initiation of the culture.
Skin equivalents were harvested and embedded in the OCT compound or fixed with 10% neutral buffered formalin for paraffin embedding for further histological, immunofluorescence and western blot analyses.
Immunofluorescence staining
Frozen sections were prepared for staining of keratin 10 (K10), keratin 14 (K14), loricrin, claudin 1, α6 integrin, type XVII collagen, Bullous pemphigoid antigen 1 (BP230/BPAG1) and laminins by fixing with acetone at -20°C for 10 min or with methanol:acetone (1:1 v/v) at room temperature, and incubating with primary antibodies overnight at 4°C. For staining of desmoglein 1 (Dsg1) and desmoglein 3 (Dsg3), nonfixed frozen sections were incubated with primary antibodies recognizing canine Dsg1 and Dsg3 for 1 h at room temperature. Filaggrin monomers were stained in paraffin-embedded sections. The primary antibodies used are listed in Table 1. Cross-reactivities of anti-K10, anti-claudin 1 and anti-laminin antibodies against canine antigens have been described by the manufacturers. Immunoreactivities of human pemphigus foliaceus sera and AK15 anti-Dsg3 monoclonal antibody to canine Dsg1 and Dsg3, respectively, were determined by immunoprecipitation using baculovirus-expressed recombinant canine Dsg115 and Dsg3 (K. Nishifuji, unpublished observation). The amino acid sequence of the loricrin peptide used as an immunogen to generate the anti-loricrin antibody in this study was mostly conserved according to the predicted amino acid sequence of canine loricrin (93%; Genbank accession no. XP 864440.2). Sections were subsequently incubated with fluorescence-conjugated secondary antibodies and Hoechst 33258 (Invitrogen) for 1 h at room temperature. All sections were subjected to microscopic analysis with a fluorescence microscope (TE2000-U; Nikon, Tokyo, Japan).
Table 1. Antibodies
Western blot analysis
Total cell lysates from the skin equivalent were applied to SDS-PAGE and blotted to polyvinylidene fluoride membranes. The bands corresponding to K14, filaggrin monomers, claudin 1 and Dsgs were visualized using the antibodies listed in Table 1.
RT-PCR analysis
Total RNA was purified from the canine skin equivalents using a RNeasy Mini Kit (Qiagen, Hilden, Germany) and used to synthesize complementary DNA using a PrimeScript RT reagent kit (Takara Bio, Shiga, Japan) according to the manufacturer’s protocol. The PCR was conducted using the AmpliTaq Gold 360 Master Mix (Applied Biosystems, Foster City, CA, USA). Reactions were performed using the following cycling conditions: 30 cycles of 30 s at 95°C, 30 s at 60°C and 60 s at 72°C. The primer sets used are shown in Table 2.
Table 2. Primer sequences for RT-PCR
| Gene | Primer sequences |
| Type XVII collagen | Forward: 5′-AAGGAGCCAAACACGAGAGA-3′ |
| Reverse: 5′-GGACTCACACTTGCTGATCG-3′ | |
| β4 Integrin | Forward: 5′-TGGACAACCTCAAGCAGATG-3′ |
| Reverse: 5′-CAGGCCTCATGTCTGTCTGA-3′ | |
| Laminin a3 | Forward: 5′-AGTTGAGGTTCACCGGTTTG-3′ |
| Reverse: 5′-GGGTGACTTGCAGGCTATGT-3′ | |
| Laminin β3 | Forward: 5′-GATTGACCAAGCCTGAGACC-3′ |
| Reverse: 5′-CCCGAAGATGAAACCACATT-3′ | |
| Laminin γ2 | Forward: 5′-GAAACCCAGCAGCTCTTACG-3′ |
| Reverse: 5′-AAAAGTGGCATTGCCCATAC-3′ | |
| Keratin 10 | Forward: 5′-TTGAGACGCACTGTTCAAGG-3′ |
| Reverse: 5′-ACGCAGTAGCGACCTTCTGT-3′ | |
| Keratin 14 | Forward: 5′-GAGATGCGTGACCAGTACGA-3′ |
| Reverse: 5′-GCAATTCGCTATTAGAGACCAC-3′ | |
| Keratin 1 | Forward: 5′-CAACCAGAGCCTTCTCCAAC-3′ |
| Reverse: 5′-TCCGAGTGGAGGTGTCTACC-3′ | |
| Keratin 5 | Forward: 5′-GACGCTGCCTACATGAACAA-3′ |
| Reverse: 5′-TCATACTGGGCCTTGACCTC-3′ |
Results
Canine bulge cell-enriched hair follicle keratinocytes undergo normal differentiation of interfollicular epidermal keratinocytes in a skin equivalent
Canine bulge-enriched keratinocytes were seeded onto canine dermal equivalents and cultured at the air-liquid interface. Reconstituted skin equivalents were harvested on days 7-14 and subjected to histological analysis. Keratinocytes in skin equivalents became stratified into three or four layers covered with a cornified layer on days 10-14, as shown by haematoxylin and eosin staining (Figure 1). Basophilic cytoplasmic granules resembling keratohyalin granules were observed in the superficial layer of the reconstituted epithelium.
Figure 1. Histological findings for canine skin equivalent. Canine bulge cell-enriched keratinocytes were placed on a canine dermal equivalent and cultured for 14 days. The keratinocytes formed three to four layers of stratified epidermis. Cytoplasmic granules resembling keratohyalin granules were visible immediately beneath the stratum corneum. Mayer’s haematoxylin and eosin stain. Scale bar represents 50 μm.
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