Letter to the Editor
exogenous non-controlled factors over a prolonged period of time. As a consequence, the sample to sample variability is significantly reduced compared with human samples and they represent reproducible standard models for proof of concept and interlaboratory comparative studies of molecular activity, permeability and safety.
Acknowledgements The research conducted at the GCAPS was supported by the French National Research Agency ANR-12-JSV5-0003 CARE. The work conducted at Focas Research Institute was supported by the NBIP, Ireland, and by Science Foundation Ireland under Grant Number 11/PI/08. Additional financial support for the collaboration has been awarded
under the Ulysses exchange program 2012–2013 funded by the ‘ministeres des Affaires etrangeres (MAE) et de l’Enseignement superieur et de la Recherche (MESR)’ in France and the Irish Research Council. Ali Tfayli, Franck Bonnier, Hugh. J. Byrne and Arlette Baillet-Guffroy designed the research study. Zeineb Farhane, Ali Tfayli, Franck Bonnier and Danielle Libong performed the research. Ali Tfayli, Franck Bonnier and Danielle Libong contributed essential reagents or tools. Ali Tfayli and Franck Bonnier analysed the data. Ali Tfayli, Franck Bonnier, Zeineb Farhane, Hugh. J. Byrne and Arlette Baillet-Guffroy wrote the paper.
Conflict of interests The authors have declared no conflicting interests.
References
1 Barbotteau Y, Gontier E, Barberet P et al. Nucl Instrum Methods Phys Res, Sect B 2005: 231: 286–291. 2 Netzlaff F, Lehr C M, Wertz P W et al. Eur J Pharm Biopharm 2005: 60: 167–178. 3 EU, COM (2013) 135 final, Communication from the commission to the European Parliament and the council in, vol. 135 2013. 4 Ajani G, Sato N, Mack J A et al. Exp Cell Res 2007: 313: 3005–3015. 5 Brohem C A, Cardeal L B, Tiago M et al. Pigment Cell Melanoma Res 2010: 24: 35–50. 6 Boyce S T, Warden G D. Am J Surg 2002: 183: 445–456. 7 El Ghalbzouri A, Siamari R, Willemze R et al. Toxicol In Vitro 2008: 22: 1311–1320. 8 Ponec M, Boelsma E, Weerheim A et al. Int J Pharm 2000: 203: 211–225. 9 Ponec M. Adv Drug Deliv Rev 2002: 54(Suppl 1): S19–S30. 10 Ponec M, Boelsma E, Gibbs S et al. Skin Pharmacol Appl Skin Physiol 2002: 15(Suppl 1): 4– 17. 11 Kojima H, Katoh M, Shinoda S et al. J Appl Toxicol 2013: doi: 10.1002/jat.2937. [Epub ahead of print]
12 Alepee N, Grandidier M H, Cotovio J. Toxicol In Vitro 2014: 28: 131–145. 13 Boelsma E, Gibbs S, Faller C et al. Acta Derm Venereol 2000: 80: 82–88. 14 Elbayed K, Berl V, Debeuckelaere C et al. Chem Res Toxicol, 2013: 26: 136–145. 15 Garcia N, Doucet O, Bayer M et al. Int J Cosmet Sci 2002: 24: 25–34. 16 Tfayli A, Piot O, Draux F et al. Biopolymers 2007: 87: 261–274. 17 Netzlaff F, Kaca M, Bock U et al. Eur J Pharm Biopharm 2007: 66: 127–134. 18 Lotte C, Patouillet C, Zanini M et al. Skin Pharmacol Appl Skin Physiol 2002: 15(Suppl 1): 18–30. 19 Schreiber S, Mahmoud A, Vuia A et al. Toxicol In Vitro 2005: 19: 813–822. 20 Kuchler S, Struver K, Friess W. Expert Opin Drug Metab Toxicol 2013: 9: 1255–1263. 21 Rozman B, Gasperlin M, Tinois-Tessoneaud E et al. Eur J Pharm Biopharm 2009: 72: 69–75. 22 Vyumvuhore R, Tfayli A, Duplan H et al. J Raman Spectrosc 2013: 44: 1077–1083. 23 Plasencia I, Norlen L, Bagatolli L A. Biophys J 2007: 93: 3142–3155. 24 Mack Correa M C, Mao G, Saad P et al. Exp Dermatol 2014: 23: 39–44.
DOI: 10.1111/exd.12424 www.wileyonlinelibrary.com/journal/EXD
25 Marcott C, Lo M, Kjoller K et al. Exp Dermatol 2013: 22: 419–421. 26 Tfayli A, Jamal D, Vyumvuhore R et al. Analyst 2013: 138: 6582–6588. 27 Guillard E, Tfayli A, Manfait M et al. Anal Bioanal Chem 2011: 399: 1201–1213. 28 Tfayli A, Guillard E, Manfait M et al. Anal Bioanal Chem 2010: 397: 1281–1296. 29 Tfayli A, Guillard E, Manfait M et al. Analyst 2012: 137: 5002–5010. 30 Tfayli A, Guillard E, Manfait M et al. Eur J Dermatol 2012: 22: 36–41.
Supporting Information Additional Supporting Information may be found in the online version of this article: Figure S1. Lipid profiles of EpiDerm model (red) and a human model (blue). Figure S2. Surface of Epiderm skin model. Figure S3. NCLS mapping on human SC representing the distribution of 1: keratin. 2: cholesterol. 3: ceramides and fatty acids. Table S1. Raman descriptors of Human SC and different zones of artificial SC. Values are the mean of 9 spectra. Data S1. Supplementary information.
Letter to the Editor
A model system to analyse the ability of human keratinocytes to form hair follicles Rajesh L. Thangapazham, Peter Klover, Shaowei Li, Ji-an Wang, Leonard Sperling and Thomas N. Darling Department of Dermatology, Uniformed Services University of the Health Sciences, Bethesda, MD, USA Correspondence: Thomas N. Darling, MD, PhD or Rajesh L. Thangapazham, PhD, Department of Dermatology, Uniformed Services University of Health Sciences, 4301 Jones Bridge Rd, Bethesda, MD 20814, USA, Tel.: 301-295-3820, Fax: 301-295-3150, e-mails: [email protected] or [email protected] Abstract: Earlier studies showed that dermal cells lose trichogenic capacity with passage, but studies on the effect of keratinocyte passage on human hair follicle neogenesis and graft quality have been hampered by the lack of a suitable model system. We recently documented human hair follicle neogenesis in grafted dermal-epidermal composites, and in the present study, we determined the effects of keratinocyte passage on hair
ª 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd Experimental Dermatology, 2014, 23, 424–448
follicle neogenesis. Dermal equivalents were made with cultured human dermal papilla cells and were overlaid with either primary or passaged human keratinocytes to form dermalepidermal composites; these were then grafted onto immunodeficient mice. Superior hair follicle neogenesis was observed using early keratinocyte cultures. Characteristics such as formation of hair shafts and sebaceous glands, presence of
443
Letter to the Editor
hair follicles with features of anagen or telogen follicles, and reproducible hair and skin function parameters make this model a tool to study human hair follicle neogenesis and development.
Key words: dermal papilla – hair follicle – keratinocyte – neogenesis – passage
Accepted for publication 21 April 2014
Abbreviations: DECdermal-epidermal composites; HFhair follicle; DPdermal papilla.
Background Skin substitutes currently in preclinical development show promise as completely autologous or combination of autologous/allogenic products (1–5), but methodologies are needed to endow skin substitutes with the potential to develop normal hair follicles (HFs) (4,6). HFs in skin substitutes may secrete factors to guide nerve migration to promote sensory recovery (7), enhance the secretion of antimicrobial or host defense peptides to reduce growth of micro-organisms (8,9), and provide normal appearance to the regenerated tissue. The lack of model systems to study human HF neogenesis in skin substitutes and the difficulty in maintaining the native properties of the isolated human cells are impeding the development of skin substitutes with appendages. Murine or human/mouse chimeric HF neogenesis experimental systems are currently used to evaluate mesenchymal cells for trichogenicity and epithelial cells for enhancing or supporting the neogenesis process (10). Important advances, such as the ability to counteract the decreased trichogenicity of dermal papilla (DP) cells passaged in vitro (11,12) by culturing as spheroids (13), have been made mostly using rodent cells in chimeric HF neogenesis models. The feasibility of modelling human HF neogenesis was evident when cultured DP cells induced HFs when primed by growing them in three-dimensional culture conditions
(a)
(14–16). Successful HF neogenesis was demonstrated using human keratinocytes and murine DP cells, whereas grafts with human only components in those studies failed to induce appendages (17,18). Current in vitro approaches, although promising for study of hair follicle neogenesis, have chimeric or incomplete HFs (19–22). We recently reported de novo human HF neogenesis in a xenograft model using dermal-epidermal composites (DECs) constructed with human cells and grafted them onto nude mice (23).
Questions addressed Does keratinocyte passage affect HF neogenesis in grafted DECs or alter graft characteristics such as size or skin barrier function? Do we observe different stages of HF in grafted DECs?
Experimental design Detailed materials and methods are provided in Data S1.
Results To study the effect of keratinocyte passage, we grafted DECs composed of the same batch of cultured DP cells combined with either primary (P0) or passaged keratinocytes. Eight weeks after grafting all the groups showed HF neogenesis, however, with varying degree (Fig. 1a, b and c). All of the grafts using P0 keratinocytes (6/6) had HFs; 83% of grafts using passage 1 keratinocytes (P1, 5/ 6) had HFs; and only 71% of grafts using passage 3 keratinocytes
(d) (f)
(g)
(b) (e) (h)
(c)
Figure 1. Characteristics of grafted dermal–epidermal composite hair follicle model. Representative images showing human hair follicle neogenesis in grafted dermal– epidermal composites made with cultured dermal papilla cells and with either primary or passaged keratinocytes 8 weeks after grafting. Primary (a) or P1 (b) keratinocytes form more hair follicles than P3 (c) keratinocytes. (d) Representative graft with hair shafts 12 weeks after grafting. (e) Selected area magnified to show pigmented hair shafts clearly. After 15 weeks, the grafts containing DP cells grown in monolayers plus those grown as spheroids had catagen/telogen hair follicles, confirmed by club-like appearance and spiky keratin fibres (f), existence of secondary hair germ with adjacent hair papilla (g, arrow) and presence of a cornified club (h, negative for toluidine blue staining, arrow). Scale bars: a, b, and c 320 lm, f 35 lm, g 65 lm and h 130 lm.
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ª 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd Experimental Dermatology, 2014, 23, 424–448
Letter to the Editor
Table 1. Graft characteristics and functional alterations in grafted dermal– epidermal composites constructed with different keratinocyte cultures Keratinocyte passage Graft characteristics and skin barrier functions1,2
Primary (P0)
Graft diameter in mm, 4 weeks3 Graft diameter in mm4 Hair follicles/mm of epidermis4 Epidermal thickness in lm4 Follicular area/dermal area in percentage4 Hair follicle diameter in lm5 Shaft diameter in lm6 TEWL in g/m2h7 Water content7 Surface hydration8
6.0 5.8 0.9 127 16
1.4 1.2 0.5 14 12
6.5 5.5 0.8 117 12
1.0 1.1 0.6 13 9
6.4 4.4 0.2 89 4
1.9 1.6 0.2* 12*** 3
250 45 27 24 3
38** 13 8 6 4
144 21 29 21 7
27 3* 6 8 2
107 24 34 18 4
22 8 15 9 3
Passage 1 (P1)
Passage 3 (P3)
1
Measured at 8 weeks except when mentioned otherwise. Results expressed as Mean SD. 3 P0, n = 8; P1, n = 7; P3, n = 8. 4 P0, n = 6; P1, n = 6; P3, n = 7. 5 P0, n = 5; P1, n = 5; P3, n = 3. 6 P0, n = 3; P1, n = 3 P3, n = 2. 7 P0, n = 6; P1, n = 6; P3, n = 5. 8 P0, n = 3; P1, n = 5; P3, n = 4. * Significantly less than primary keratinocytes, P < 0.05. *** Significantly less than primary and passage 1 keratinocytes, P < 0.005. ** Significantly greater than passaged keratinocytes, P < 0.02. 2
(P3, 5/7) had HFs. Grafts with P3 keratinocytes had significantly thinner epidermal layer than grafts made with either P0 or P1 keratinocytes. Grafts using P3 keratinocytes had a significantly lower HF density than grafts using P0 keratinocytes (Table 1). HF diameter and shaft diameter using P0 keratinocytes were greater than those observed in composites using P3 and P1 keratinocytes, respectively (Table 1). Follicular area per dermal area was not significantly different between the groups. The hair follicle length using early passage keratinocytes, measured from bulb to infundibulum in the follicles sectioned along their entire length, was 620 120 lm (n = 5). To study and compare the functional alterations of the grafted DECs with different keratinocytes, we measured graft diameter, transepidermal water loss, skin water content and surface hydration. Grafts made with lower passage keratinocytes trended towards lower transepidermal water loss rate and higher water content (Table 1), but these results were not statistically significant. Hair follicles in 6 of 19 grafts with HFs harvested at 8 weeks were associated with sebaceous glands that secrete an antimicrobial peptide, cathelicidin (Figure S2a, b). Arrector pili were not observed in any of the grafts. After 10 weeks, we observed longer pigmented hair shafts (Fig. 1d, e). We verified human origin of
References
1 Pham C, Greenwood J, Cleland H et al. Burns 2007: 33: 946–957. 2 Guerret S, Govignon E, Hartmann D J et al. Wound Repair Regen 2003: 11: 35–45. 3 Centanni J M, Straseski J A, Wicks A et al. Ann Surg 2011: 253: 672–683. 4 Sriwiriyanont P, Lynch K A, Maier E A et al. Exp Dermatol 2012: 21: 783–785. 5 Killat J, Reimers K, Choi C Y et al. Int J Mol Sci 2013: 14: 14460–14474. 6 Ohyama M, Veraitch O. J Dermatol Sci 2013: 70: 78–87.
the HFs using human-specific ALU probe (Figure S1a) and human-specific COX IV staining (Figure S1b). All the HFs that we have so far observed in grafts harvested at 8 weeks were in anagen, whereas HFs in 3 grafts harvested after 10 weeks were classified as anagen [11 of 26 HFs counted (42%)], catagen [one (4%)] or telogen [14 (54%)]. Telogen HFs were observed as early as 10 weeks and had scarce Ki-67 positive cells, whereas anagen HFs had larger population of Ki-67 positive cells in the matrix (Figure S1c, d). The presence of telogen HFs in 15-week grafts (Fig. 1f) was further substantiated by their club-like appearance, spiky keratin fibres in the inner root sheath (Fig. 1f), a secondary hair germ with adjacent hair papilla (Fig. 1g) and the presence of a cornified club that stains negative with toluidine blue (Fig. 1h).
Conclusions We observed enhanced HF neogenesis with early passage keratinocytes which may be due to the presence of more progenitor cells in the early passage keratinocytes. Previous studies have demonstrated the abundance of epidermal progenitor cells like CD90 (+) (24) and a6integrinbriCD71dim (25) in primary keratinocyte cultures and their dearth in later passage. These cultures enriched for progenitor cells have high colony-forming capability (24,25) and form a significantly thicker epithelium in an organotypic skin equivalent system (25). Hence, we hypothesize that early keratinocyte cultures or factors that retain the precursor cells are pivotal for HF neogenesis and subsequently for developing skin appendages with hair follicles. The observation of hair follicles classified as anagen, catagen or telogen at 10 weeks or later suggests that this model system also permits the evaluation of transit through these phases. Further studies are required to determine whether anagen can be reinduced in these DECs. Overall, our results suggest that this model system can be utilized to evaluate genes and cells critical for human HF neogenesis and development or to optimize autologous tissue engineered skin substitutes for clinical applications.
Acknowledgements This work was supported by a grant from the Defense Medical Research and Development Program to Thomas Darling, M.D. Ph.D and National Institutes of Health Skin Disease Research Center grant 5-P30-AR-057217.
Author contributions TND and RLT conceived the study, designed the experiments and wrote the manuscript. RLT, PK, JW and SL performed the experiments. LS analysed the hair follicle morphology and stages. TND supervised the research.
Conflict of interest The authors have declared no conflicting interests.
7 Gagnon V, Larouche D, Parenteau-Bareil R et al. J Invest Dermatol 2011: 131: 1375–1378. 8 Chronnell C M, Ghali L R, Ali R S et al. J Invest Dermatol 2001: 117: 1120–1125. 9 Gibson A L, Thomas-Virnig C L, Centanni J M et al. Wound Repair Regen 2012: 20: 414– 424. 10 Yang C C, Cotsarelis G. J Dermatol Sci 2010: 57: 2–11. 11 Horne K A, Jahoda C A, Oliver R F et al. J Embryol Exp Morphol 1986: 97: 111–124. 12 Soma T, Fujiwara S, Shirakata Y et al. Exp Dermatol 2012: 21: 307–309.
ª 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd Experimental Dermatology, 2014, 23, 424–448
13 Osada A, Iwabuchi T, Kishimoto J et al. Tissue Eng 2007: 13: 975–982. 14 Higgins C A, Richardson G D, Ferdinando D et al. Exp Dermatol 2010: 19: 546–548. 15 Kang B M, Kwack M H, Kim M K et al. J Invest Dermatol 2012: 132: 237–239. 16 Higgins C A, Chen J C, Cerise J E et al. Proc Natl Acad Sci U S A 2013: 110: 19679–19688. 17 Ehama R, Ishimatsu-Tsuji Y, Iriyama S et al. J Invest Dermatol 2007: 127: 2106–2115. 18 Sriwiriyanont P, Lynch K A, McFarland K L et al. PLoS ONE 2013: 8: e65664.
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19 Krugluger W, Rohrbacher W, Laciak K et al. Exp Dermatol 2005: 14: 580–585. 20 Qiao J, Turetsky A, Kemp P et al. Regen Med 2008: 3: 683–692. T, Mescalchin A et al. J Invest 21 Havlickova B, Bıro Dermatol 2009: 129: 972–983. 22 Lindner G, Horland R, Wagner I et al. J Biotechnol 2011: 152: 108–112. 23 Thangapazham R L, Klover P, Wang J A et al. J Invest Dermatol 2013: 134: 538–540. 24 Nakamura Y, Muguruma Y, Yahata T et al. Br J Dermatol 2006: 154: 1062–1070. 25 Fujimori Y, Izumi K, Feinberg S E et al. J Dermatol Sci 2009: 56: 181–187.
Supporting Information Additional Supporting Information may be found in the online version of this article:
Data S1. Methods. Figure S1. (a) Fluorescence in situ hybridization (FISH) analysis using a human-specific ALU probe (green) showing hybridization to nuclei in epithelial and dermal cells, including dermal papilla and dermal sheath. (b) H&E stained section of dermal–epidermal composite harvested at 10 weeks shows junction of graft and host mouse skin. (c) Serial section of (b) stained with the antibody specific for human but not mouse COX IV confirms junction of graft and host mouse skin (dotted line). Human cells are immunoreactive in graft epithelium and dermis as well as in the de novo regenerated follicular epithelium (arrow), whereas adjacent mouse dermal cells, and follicular (arrow heads) and interfollicular epithelium stained negative. (d) Telogen HF from serial section of (c) shows no Ki-67 positive cells consistent with telogen
DOI: 10.1111/exd.12425 www.wileyonlinelibrary.com/journal/EXD
stage of hair follicle (arrow). (e) An anagen hair follicle (arrow) with dermal papilla from the same section as the telogen hair follicle in (d) shows dense Ki-67 reactivity in matrix as expected. Scale bars: a 50 lm, b and c 320 lm, and d and e 35 lm. Figure S2. (a) Representative H&E stained section of graft showing hair follicle inner and outer root sheath and sebaceous gland. (b) Sebaceous gland was highly immunoreactive to an antibody for cathelicidin, an antimicrobial peptide. No staining was detected in glandular structure or the epithelium of the sebaceous gland in a serial section stained with an irrelevant IgG, which served as a negative control (image not shown). Scale bars: a and b 35 lm.
Letter to the Editor
Similar appearance, different mechanisms: xerosis in HIV, atopic dermatitis and ageing € ndermann1, Diedrich A. Schmidt1†, Anja Potthoff4, Meike Mischo1*, Laura B. von Kobyletzki2,3*, Erik Bru 4,5 1 Norbert H. Brockmeyer and Martina Havenith 1
Physical Chemistry II, Ruhr-Universit€at Bochum, Bochum, Germany; 2Department of Dermatology, Institute of Clinical Research in Malm€ o, Lund University, Sk ane University Hospital, Malm€ o, Sweden; 3Department of Public Health Sciences, Karlstad University, Karlstad, Sweden; 4Department of Dermatology, Ruhr-University Bochum, Bochum, Germany and 5Competence Network for HIV/Aids, Ruhr-University Bochum, Bochum, Germany; Correspondence: Martina Havenith, Physical Chemistry II, Universit€atsstr. 150, 44801 Bochum, Germany, Tel.: +49-234-32-28249, Fax: +49-234-32-14183, e-mail: [email protected] † Present Address: Department of Physics, North Carolina A&T State University, Greensboro, NC, USA and The Joint School of Nanoscience & Nanoengineering (A&T and UNCG), Greensboro, NC, USA *These authors contributed equally to this work. Abstract: Xerosis is one of the most common dermatologic disorders occurring in the elderly and in patients with atopic dermatitis (AD) and human immunodeficiency virus (HIV) infection. Xerosis has been linked to an impaired skin barrier function of the stratum corneum. Using Raman microspectroscopy, we concentrated on deeper skin layers, viable epidermis and dermis of 47 volunteers and associated molecular alterations to the evolution of xerosis and the skin barrier, for example, lipid, water and antioxidant content. A decrease in lipids within the viable epidermis is found for elderly and HIV-patients. Lipid and water values of AD patients and their healthy reference
group are similar. Decreases in lipids and simultaneous increases in water are found in the dermis for HIV and AD patients in comparison to their healthy reference groups. Excessive levels of epidermal carotenoids, mainly lycopene, in HIV-patients were found potentially leading to adverse effects such as premature skin ageing.
Background
aged humans and mice (10) has been correlated with impaired SC lipid processing (3). The underlying mechanisms of AD affect lamellar bodies (LB) maturation, LB secretion and lipid processing (3). In elderly human and mouse skin, LBs are formed, but are void of lipids (11,12); however, lipid synthesis is unchanged from young to moderately aged murine skin (10). Transepidermal water loss (TEWL), a combined effect of content and transport, is a measure to assess water metabolism, which can increase for AD at affected sites, but not at unaffected sites (13). Basal TEWL levels decrease from elderly to younger people,
Xerosis, a common dermatological disorder in the elderly (1), a signature of atopic dermatitis (AD), and co-morbidity with human immunodeficiency virus (HIV) infection (2–4) has been associated with an impaired skin barrier function in the stratum corneum (SC) due to lipid lamellae modifications and regulation of hydration (5–7). However, deeper skin layers play critical roles: the dermis delivers nutrients to the viable epidermis (VE), which produces components for the SC. Skin appearance due to ageing is often associated to free radicals (8,9). Increased pH in the SC of
446
Key words: atopic dermatitis – HIV – molecular composition – Raman – xerosis
Accepted for publication 21 April 2014
ª 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd Experimental Dermatology, 2014, 23, 424–448
exogenous non-controlled factors over a prolonged period of time. As a consequence, the sample to sample variability is significantly reduced compared with human samples and they represent reproducible standard models for proof of concept and interlaboratory comparative studies of molecular activity, permeability and safety.
Acknowledgements The research conducted at the GCAPS was supported by the French National Research Agency ANR-12-JSV5-0003 CARE. The work conducted at Focas Research Institute was supported by the NBIP, Ireland, and by Science Foundation Ireland under Grant Number 11/PI/08. Additional financial support for the collaboration has been awarded
under the Ulysses exchange program 2012–2013 funded by the ‘ministeres des Affaires etrangeres (MAE) et de l’Enseignement superieur et de la Recherche (MESR)’ in France and the Irish Research Council. Ali Tfayli, Franck Bonnier, Hugh. J. Byrne and Arlette Baillet-Guffroy designed the research study. Zeineb Farhane, Ali Tfayli, Franck Bonnier and Danielle Libong performed the research. Ali Tfayli, Franck Bonnier and Danielle Libong contributed essential reagents or tools. Ali Tfayli and Franck Bonnier analysed the data. Ali Tfayli, Franck Bonnier, Zeineb Farhane, Hugh. J. Byrne and Arlette Baillet-Guffroy wrote the paper.
Conflict of interests The authors have declared no conflicting interests.
References
1 Barbotteau Y, Gontier E, Barberet P et al. Nucl Instrum Methods Phys Res, Sect B 2005: 231: 286–291. 2 Netzlaff F, Lehr C M, Wertz P W et al. Eur J Pharm Biopharm 2005: 60: 167–178. 3 EU, COM (2013) 135 final, Communication from the commission to the European Parliament and the council in, vol. 135 2013. 4 Ajani G, Sato N, Mack J A et al. Exp Cell Res 2007: 313: 3005–3015. 5 Brohem C A, Cardeal L B, Tiago M et al. Pigment Cell Melanoma Res 2010: 24: 35–50. 6 Boyce S T, Warden G D. Am J Surg 2002: 183: 445–456. 7 El Ghalbzouri A, Siamari R, Willemze R et al. Toxicol In Vitro 2008: 22: 1311–1320. 8 Ponec M, Boelsma E, Weerheim A et al. Int J Pharm 2000: 203: 211–225. 9 Ponec M. Adv Drug Deliv Rev 2002: 54(Suppl 1): S19–S30. 10 Ponec M, Boelsma E, Gibbs S et al. Skin Pharmacol Appl Skin Physiol 2002: 15(Suppl 1): 4– 17. 11 Kojima H, Katoh M, Shinoda S et al. J Appl Toxicol 2013: doi: 10.1002/jat.2937. [Epub ahead of print]
12 Alepee N, Grandidier M H, Cotovio J. Toxicol In Vitro 2014: 28: 131–145. 13 Boelsma E, Gibbs S, Faller C et al. Acta Derm Venereol 2000: 80: 82–88. 14 Elbayed K, Berl V, Debeuckelaere C et al. Chem Res Toxicol, 2013: 26: 136–145. 15 Garcia N, Doucet O, Bayer M et al. Int J Cosmet Sci 2002: 24: 25–34. 16 Tfayli A, Piot O, Draux F et al. Biopolymers 2007: 87: 261–274. 17 Netzlaff F, Kaca M, Bock U et al. Eur J Pharm Biopharm 2007: 66: 127–134. 18 Lotte C, Patouillet C, Zanini M et al. Skin Pharmacol Appl Skin Physiol 2002: 15(Suppl 1): 18–30. 19 Schreiber S, Mahmoud A, Vuia A et al. Toxicol In Vitro 2005: 19: 813–822. 20 Kuchler S, Struver K, Friess W. Expert Opin Drug Metab Toxicol 2013: 9: 1255–1263. 21 Rozman B, Gasperlin M, Tinois-Tessoneaud E et al. Eur J Pharm Biopharm 2009: 72: 69–75. 22 Vyumvuhore R, Tfayli A, Duplan H et al. J Raman Spectrosc 2013: 44: 1077–1083. 23 Plasencia I, Norlen L, Bagatolli L A. Biophys J 2007: 93: 3142–3155. 24 Mack Correa M C, Mao G, Saad P et al. Exp Dermatol 2014: 23: 39–44.
DOI: 10.1111/exd.12424 www.wileyonlinelibrary.com/journal/EXD
25 Marcott C, Lo M, Kjoller K et al. Exp Dermatol 2013: 22: 419–421. 26 Tfayli A, Jamal D, Vyumvuhore R et al. Analyst 2013: 138: 6582–6588. 27 Guillard E, Tfayli A, Manfait M et al. Anal Bioanal Chem 2011: 399: 1201–1213. 28 Tfayli A, Guillard E, Manfait M et al. Anal Bioanal Chem 2010: 397: 1281–1296. 29 Tfayli A, Guillard E, Manfait M et al. Analyst 2012: 137: 5002–5010. 30 Tfayli A, Guillard E, Manfait M et al. Eur J Dermatol 2012: 22: 36–41.
Supporting Information Additional Supporting Information may be found in the online version of this article: Figure S1. Lipid profiles of EpiDerm model (red) and a human model (blue). Figure S2. Surface of Epiderm skin model. Figure S3. NCLS mapping on human SC representing the distribution of 1: keratin. 2: cholesterol. 3: ceramides and fatty acids. Table S1. Raman descriptors of Human SC and different zones of artificial SC. Values are the mean of 9 spectra. Data S1. Supplementary information.
Letter to the Editor
A model system to analyse the ability of human keratinocytes to form hair follicles Rajesh L. Thangapazham, Peter Klover, Shaowei Li, Ji-an Wang, Leonard Sperling and Thomas N. Darling Department of Dermatology, Uniformed Services University of the Health Sciences, Bethesda, MD, USA Correspondence: Thomas N. Darling, MD, PhD or Rajesh L. Thangapazham, PhD, Department of Dermatology, Uniformed Services University of Health Sciences, 4301 Jones Bridge Rd, Bethesda, MD 20814, USA, Tel.: 301-295-3820, Fax: 301-295-3150, e-mails: [email protected] or [email protected] Abstract: Earlier studies showed that dermal cells lose trichogenic capacity with passage, but studies on the effect of keratinocyte passage on human hair follicle neogenesis and graft quality have been hampered by the lack of a suitable model system. We recently documented human hair follicle neogenesis in grafted dermal-epidermal composites, and in the present study, we determined the effects of keratinocyte passage on hair
ª 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd Experimental Dermatology, 2014, 23, 424–448
follicle neogenesis. Dermal equivalents were made with cultured human dermal papilla cells and were overlaid with either primary or passaged human keratinocytes to form dermalepidermal composites; these were then grafted onto immunodeficient mice. Superior hair follicle neogenesis was observed using early keratinocyte cultures. Characteristics such as formation of hair shafts and sebaceous glands, presence of
443
Letter to the Editor
hair follicles with features of anagen or telogen follicles, and reproducible hair and skin function parameters make this model a tool to study human hair follicle neogenesis and development.
Key words: dermal papilla – hair follicle – keratinocyte – neogenesis – passage
Accepted for publication 21 April 2014
Abbreviations: DECdermal-epidermal composites; HFhair follicle; DPdermal papilla.
Background Skin substitutes currently in preclinical development show promise as completely autologous or combination of autologous/allogenic products (1–5), but methodologies are needed to endow skin substitutes with the potential to develop normal hair follicles (HFs) (4,6). HFs in skin substitutes may secrete factors to guide nerve migration to promote sensory recovery (7), enhance the secretion of antimicrobial or host defense peptides to reduce growth of micro-organisms (8,9), and provide normal appearance to the regenerated tissue. The lack of model systems to study human HF neogenesis in skin substitutes and the difficulty in maintaining the native properties of the isolated human cells are impeding the development of skin substitutes with appendages. Murine or human/mouse chimeric HF neogenesis experimental systems are currently used to evaluate mesenchymal cells for trichogenicity and epithelial cells for enhancing or supporting the neogenesis process (10). Important advances, such as the ability to counteract the decreased trichogenicity of dermal papilla (DP) cells passaged in vitro (11,12) by culturing as spheroids (13), have been made mostly using rodent cells in chimeric HF neogenesis models. The feasibility of modelling human HF neogenesis was evident when cultured DP cells induced HFs when primed by growing them in three-dimensional culture conditions
(a)
(14–16). Successful HF neogenesis was demonstrated using human keratinocytes and murine DP cells, whereas grafts with human only components in those studies failed to induce appendages (17,18). Current in vitro approaches, although promising for study of hair follicle neogenesis, have chimeric or incomplete HFs (19–22). We recently reported de novo human HF neogenesis in a xenograft model using dermal-epidermal composites (DECs) constructed with human cells and grafted them onto nude mice (23).
Questions addressed Does keratinocyte passage affect HF neogenesis in grafted DECs or alter graft characteristics such as size or skin barrier function? Do we observe different stages of HF in grafted DECs?
Experimental design Detailed materials and methods are provided in Data S1.
Results To study the effect of keratinocyte passage, we grafted DECs composed of the same batch of cultured DP cells combined with either primary (P0) or passaged keratinocytes. Eight weeks after grafting all the groups showed HF neogenesis, however, with varying degree (Fig. 1a, b and c). All of the grafts using P0 keratinocytes (6/6) had HFs; 83% of grafts using passage 1 keratinocytes (P1, 5/ 6) had HFs; and only 71% of grafts using passage 3 keratinocytes
(d) (f)
(g)
(b) (e) (h)
(c)
Figure 1. Characteristics of grafted dermal–epidermal composite hair follicle model. Representative images showing human hair follicle neogenesis in grafted dermal– epidermal composites made with cultured dermal papilla cells and with either primary or passaged keratinocytes 8 weeks after grafting. Primary (a) or P1 (b) keratinocytes form more hair follicles than P3 (c) keratinocytes. (d) Representative graft with hair shafts 12 weeks after grafting. (e) Selected area magnified to show pigmented hair shafts clearly. After 15 weeks, the grafts containing DP cells grown in monolayers plus those grown as spheroids had catagen/telogen hair follicles, confirmed by club-like appearance and spiky keratin fibres (f), existence of secondary hair germ with adjacent hair papilla (g, arrow) and presence of a cornified club (h, negative for toluidine blue staining, arrow). Scale bars: a, b, and c 320 lm, f 35 lm, g 65 lm and h 130 lm.
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ª 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd Experimental Dermatology, 2014, 23, 424–448
Letter to the Editor
Table 1. Graft characteristics and functional alterations in grafted dermal– epidermal composites constructed with different keratinocyte cultures Keratinocyte passage Graft characteristics and skin barrier functions1,2
Primary (P0)
Graft diameter in mm, 4 weeks3 Graft diameter in mm4 Hair follicles/mm of epidermis4 Epidermal thickness in lm4 Follicular area/dermal area in percentage4 Hair follicle diameter in lm5 Shaft diameter in lm6 TEWL in g/m2h7 Water content7 Surface hydration8
6.0 5.8 0.9 127 16
1.4 1.2 0.5 14 12
6.5 5.5 0.8 117 12
1.0 1.1 0.6 13 9
6.4 4.4 0.2 89 4
1.9 1.6 0.2* 12*** 3
250 45 27 24 3
38** 13 8 6 4
144 21 29 21 7
27 3* 6 8 2
107 24 34 18 4
22 8 15 9 3
Passage 1 (P1)
Passage 3 (P3)
1
Measured at 8 weeks except when mentioned otherwise. Results expressed as Mean SD. 3 P0, n = 8; P1, n = 7; P3, n = 8. 4 P0, n = 6; P1, n = 6; P3, n = 7. 5 P0, n = 5; P1, n = 5; P3, n = 3. 6 P0, n = 3; P1, n = 3 P3, n = 2. 7 P0, n = 6; P1, n = 6; P3, n = 5. 8 P0, n = 3; P1, n = 5; P3, n = 4. * Significantly less than primary keratinocytes, P < 0.05. *** Significantly less than primary and passage 1 keratinocytes, P < 0.005. ** Significantly greater than passaged keratinocytes, P < 0.02. 2
(P3, 5/7) had HFs. Grafts with P3 keratinocytes had significantly thinner epidermal layer than grafts made with either P0 or P1 keratinocytes. Grafts using P3 keratinocytes had a significantly lower HF density than grafts using P0 keratinocytes (Table 1). HF diameter and shaft diameter using P0 keratinocytes were greater than those observed in composites using P3 and P1 keratinocytes, respectively (Table 1). Follicular area per dermal area was not significantly different between the groups. The hair follicle length using early passage keratinocytes, measured from bulb to infundibulum in the follicles sectioned along their entire length, was 620 120 lm (n = 5). To study and compare the functional alterations of the grafted DECs with different keratinocytes, we measured graft diameter, transepidermal water loss, skin water content and surface hydration. Grafts made with lower passage keratinocytes trended towards lower transepidermal water loss rate and higher water content (Table 1), but these results were not statistically significant. Hair follicles in 6 of 19 grafts with HFs harvested at 8 weeks were associated with sebaceous glands that secrete an antimicrobial peptide, cathelicidin (Figure S2a, b). Arrector pili were not observed in any of the grafts. After 10 weeks, we observed longer pigmented hair shafts (Fig. 1d, e). We verified human origin of
References
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the HFs using human-specific ALU probe (Figure S1a) and human-specific COX IV staining (Figure S1b). All the HFs that we have so far observed in grafts harvested at 8 weeks were in anagen, whereas HFs in 3 grafts harvested after 10 weeks were classified as anagen [11 of 26 HFs counted (42%)], catagen [one (4%)] or telogen [14 (54%)]. Telogen HFs were observed as early as 10 weeks and had scarce Ki-67 positive cells, whereas anagen HFs had larger population of Ki-67 positive cells in the matrix (Figure S1c, d). The presence of telogen HFs in 15-week grafts (Fig. 1f) was further substantiated by their club-like appearance, spiky keratin fibres in the inner root sheath (Fig. 1f), a secondary hair germ with adjacent hair papilla (Fig. 1g) and the presence of a cornified club that stains negative with toluidine blue (Fig. 1h).
Conclusions We observed enhanced HF neogenesis with early passage keratinocytes which may be due to the presence of more progenitor cells in the early passage keratinocytes. Previous studies have demonstrated the abundance of epidermal progenitor cells like CD90 (+) (24) and a6integrinbriCD71dim (25) in primary keratinocyte cultures and their dearth in later passage. These cultures enriched for progenitor cells have high colony-forming capability (24,25) and form a significantly thicker epithelium in an organotypic skin equivalent system (25). Hence, we hypothesize that early keratinocyte cultures or factors that retain the precursor cells are pivotal for HF neogenesis and subsequently for developing skin appendages with hair follicles. The observation of hair follicles classified as anagen, catagen or telogen at 10 weeks or later suggests that this model system also permits the evaluation of transit through these phases. Further studies are required to determine whether anagen can be reinduced in these DECs. Overall, our results suggest that this model system can be utilized to evaluate genes and cells critical for human HF neogenesis and development or to optimize autologous tissue engineered skin substitutes for clinical applications.
Acknowledgements This work was supported by a grant from the Defense Medical Research and Development Program to Thomas Darling, M.D. Ph.D and National Institutes of Health Skin Disease Research Center grant 5-P30-AR-057217.
Author contributions TND and RLT conceived the study, designed the experiments and wrote the manuscript. RLT, PK, JW and SL performed the experiments. LS analysed the hair follicle morphology and stages. TND supervised the research.
Conflict of interest The authors have declared no conflicting interests.
7 Gagnon V, Larouche D, Parenteau-Bareil R et al. J Invest Dermatol 2011: 131: 1375–1378. 8 Chronnell C M, Ghali L R, Ali R S et al. J Invest Dermatol 2001: 117: 1120–1125. 9 Gibson A L, Thomas-Virnig C L, Centanni J M et al. Wound Repair Regen 2012: 20: 414– 424. 10 Yang C C, Cotsarelis G. J Dermatol Sci 2010: 57: 2–11. 11 Horne K A, Jahoda C A, Oliver R F et al. J Embryol Exp Morphol 1986: 97: 111–124. 12 Soma T, Fujiwara S, Shirakata Y et al. Exp Dermatol 2012: 21: 307–309.
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13 Osada A, Iwabuchi T, Kishimoto J et al. Tissue Eng 2007: 13: 975–982. 14 Higgins C A, Richardson G D, Ferdinando D et al. Exp Dermatol 2010: 19: 546–548. 15 Kang B M, Kwack M H, Kim M K et al. J Invest Dermatol 2012: 132: 237–239. 16 Higgins C A, Chen J C, Cerise J E et al. Proc Natl Acad Sci U S A 2013: 110: 19679–19688. 17 Ehama R, Ishimatsu-Tsuji Y, Iriyama S et al. J Invest Dermatol 2007: 127: 2106–2115. 18 Sriwiriyanont P, Lynch K A, McFarland K L et al. PLoS ONE 2013: 8: e65664.
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19 Krugluger W, Rohrbacher W, Laciak K et al. Exp Dermatol 2005: 14: 580–585. 20 Qiao J, Turetsky A, Kemp P et al. Regen Med 2008: 3: 683–692. T, Mescalchin A et al. J Invest 21 Havlickova B, Bıro Dermatol 2009: 129: 972–983. 22 Lindner G, Horland R, Wagner I et al. J Biotechnol 2011: 152: 108–112. 23 Thangapazham R L, Klover P, Wang J A et al. J Invest Dermatol 2013: 134: 538–540. 24 Nakamura Y, Muguruma Y, Yahata T et al. Br J Dermatol 2006: 154: 1062–1070. 25 Fujimori Y, Izumi K, Feinberg S E et al. J Dermatol Sci 2009: 56: 181–187.
Supporting Information Additional Supporting Information may be found in the online version of this article:
Data S1. Methods. Figure S1. (a) Fluorescence in situ hybridization (FISH) analysis using a human-specific ALU probe (green) showing hybridization to nuclei in epithelial and dermal cells, including dermal papilla and dermal sheath. (b) H&E stained section of dermal–epidermal composite harvested at 10 weeks shows junction of graft and host mouse skin. (c) Serial section of (b) stained with the antibody specific for human but not mouse COX IV confirms junction of graft and host mouse skin (dotted line). Human cells are immunoreactive in graft epithelium and dermis as well as in the de novo regenerated follicular epithelium (arrow), whereas adjacent mouse dermal cells, and follicular (arrow heads) and interfollicular epithelium stained negative. (d) Telogen HF from serial section of (c) shows no Ki-67 positive cells consistent with telogen
DOI: 10.1111/exd.12425 www.wileyonlinelibrary.com/journal/EXD
stage of hair follicle (arrow). (e) An anagen hair follicle (arrow) with dermal papilla from the same section as the telogen hair follicle in (d) shows dense Ki-67 reactivity in matrix as expected. Scale bars: a 50 lm, b and c 320 lm, and d and e 35 lm. Figure S2. (a) Representative H&E stained section of graft showing hair follicle inner and outer root sheath and sebaceous gland. (b) Sebaceous gland was highly immunoreactive to an antibody for cathelicidin, an antimicrobial peptide. No staining was detected in glandular structure or the epithelium of the sebaceous gland in a serial section stained with an irrelevant IgG, which served as a negative control (image not shown). Scale bars: a and b 35 lm.
Letter to the Editor
Similar appearance, different mechanisms: xerosis in HIV, atopic dermatitis and ageing € ndermann1, Diedrich A. Schmidt1†, Anja Potthoff4, Meike Mischo1*, Laura B. von Kobyletzki2,3*, Erik Bru 4,5 1 Norbert H. Brockmeyer and Martina Havenith 1
Physical Chemistry II, Ruhr-Universit€at Bochum, Bochum, Germany; 2Department of Dermatology, Institute of Clinical Research in Malm€ o, Lund University, Sk ane University Hospital, Malm€ o, Sweden; 3Department of Public Health Sciences, Karlstad University, Karlstad, Sweden; 4Department of Dermatology, Ruhr-University Bochum, Bochum, Germany and 5Competence Network for HIV/Aids, Ruhr-University Bochum, Bochum, Germany; Correspondence: Martina Havenith, Physical Chemistry II, Universit€atsstr. 150, 44801 Bochum, Germany, Tel.: +49-234-32-28249, Fax: +49-234-32-14183, e-mail: [email protected] † Present Address: Department of Physics, North Carolina A&T State University, Greensboro, NC, USA and The Joint School of Nanoscience & Nanoengineering (A&T and UNCG), Greensboro, NC, USA *These authors contributed equally to this work. Abstract: Xerosis is one of the most common dermatologic disorders occurring in the elderly and in patients with atopic dermatitis (AD) and human immunodeficiency virus (HIV) infection. Xerosis has been linked to an impaired skin barrier function of the stratum corneum. Using Raman microspectroscopy, we concentrated on deeper skin layers, viable epidermis and dermis of 47 volunteers and associated molecular alterations to the evolution of xerosis and the skin barrier, for example, lipid, water and antioxidant content. A decrease in lipids within the viable epidermis is found for elderly and HIV-patients. Lipid and water values of AD patients and their healthy reference
group are similar. Decreases in lipids and simultaneous increases in water are found in the dermis for HIV and AD patients in comparison to their healthy reference groups. Excessive levels of epidermal carotenoids, mainly lycopene, in HIV-patients were found potentially leading to adverse effects such as premature skin ageing.
Background
aged humans and mice (10) has been correlated with impaired SC lipid processing (3). The underlying mechanisms of AD affect lamellar bodies (LB) maturation, LB secretion and lipid processing (3). In elderly human and mouse skin, LBs are formed, but are void of lipids (11,12); however, lipid synthesis is unchanged from young to moderately aged murine skin (10). Transepidermal water loss (TEWL), a combined effect of content and transport, is a measure to assess water metabolism, which can increase for AD at affected sites, but not at unaffected sites (13). Basal TEWL levels decrease from elderly to younger people,
Xerosis, a common dermatological disorder in the elderly (1), a signature of atopic dermatitis (AD), and co-morbidity with human immunodeficiency virus (HIV) infection (2–4) has been associated with an impaired skin barrier function in the stratum corneum (SC) due to lipid lamellae modifications and regulation of hydration (5–7). However, deeper skin layers play critical roles: the dermis delivers nutrients to the viable epidermis (VE), which produces components for the SC. Skin appearance due to ageing is often associated to free radicals (8,9). Increased pH in the SC of
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Key words: atopic dermatitis – HIV – molecular composition – Raman – xerosis
Accepted for publication 21 April 2014
ª 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd Experimental Dermatology, 2014, 23, 424–448
