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* Laboratoire d’Organogénèse Expérimentale/LOEX, Hôpital du Saint-Sacrement du CHA, Québec, Canada, and Department of Surgery, Laval University, Sainte-Foy, Québec, Canada; and
Departments of Biological Chemistry and Dermatology, The Johns Hopkins University School of Medicine, Baltimore, Maryland
1Correspondence: Laboratoire d’Organogénèse Expérimentale, Hôpital du St-Sacrement du CHA, 1050 Chemin Sainte-Foy, Québec, QC, Canada, G1S 4L8. E-mail: lucie.germain{at}chg.ulaval.ca
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: intermediate filaments • tissue engineering • transmission electron microscopy • rodent
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The hair follicle is a topologically complex epithelial appendage consisting of a central hair fiber surrounded by three concentric epithelial structures: the inner root sheath (IRS), the companion layer (cl) and the outer root sheath (ORS). Like other hair follicles, vibrissa cyclically grow (anagen), involute (catagen), and rest (telogen). A permanent pool of epithelial SCs ensures the production of cells to regenerate new hair at the beginning of each cycle (1
2
3
4)
. SCs have a long cell cycle, reflecting infrequent divisions. Following nucleotide analogues uptake, SCs retain the label over long time periods. Accordingly, these cells are also called label-retaining cells (LRCs). The identification of slow-cycling SCs provided the initial evidence that the bulge area houses epithelial SCs (1
, 3
, 5
, 6)
. Bulge SCs have the capacity to self-renew and give rise to hair follicles and other skin epithelia (e.g., epidermis, sebaceous glands), depending on the prevailing conditions (2
3
4
, 7
, 8)
.
The pairwise and differentiation-related regulation of type I (K9-K28; K31-K40) and II (K1-K8; K71-K86) keratin (K) genes provides a unique handle to track lineage specification and differentiation within epithelia (9)
. More than half of the 54 known type I and II keratin genes (9)
are expressed in hair follicles alone, and two specific type I keratins, K19 (5)
and K15 (10)
, occur at higher levels in a subpopulation of ORS basal cells that encompasses the bulge.
In the present study, we exploited the self-assembly approach of tissue engineering (11)
to show that the best tissue source in the vibrissa follicle for epithelial reconstruction in vitro is the bulge area. Further analysis of bulge epithelial cells with respect to proliferative status (SC identification using the LRC feature), keratin expression, and organization helped define three distinct subpopulations within vibrissa bulge. Two of these subpopulations consist of LRCs with colony-forming ability: Bulge basal keratinocytes (Bb) expressing K5, K15, K17, K19; and bulge suprabasal keratinocytes (Bs1), comprising cells strongly expressing K5/K17 correlating with an unusually high density of IFs. In mice lacking K17 (12)
, vibrissae were often absent or rudimentary, correlating with a profound disruption of the morphological features of Bs1 cells. Altogether, the findings reported point to cellular heterogeneity within the pool of epithelial SCs in the hair follicle bulge and differentiate them with regard to keratin expression, filament organization, and long-term retention of label.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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with modifications. Newborn (1 day old, P1) FVB, C57BL/6, and C57BL/6 K17–/– mice and adult (3 months old) FVB mice (Charles River, Saint-Constant, QC, Canada) were injected with 50 mg/kg body weight (BW) of 5-bromo-2'-deoxyuridine (BrdU) (Sigma Chemicals, St. Louis, MO, USA) every 12 h for 5 consecutive days. Mice were sacrificed 1 or 7 months later, and whisker pads were embedded in Tissue-Tek OCT Compound (OCT) (Bayers Canada, Etobicoke, ON, Canada). Actively proliferating cells were localized by injecting 1-month-old FVB, C57BL/6 (Charles River) and C57BL/6 K17–/– (12)
mice with 50 mg/kg BW of BrdU at 0 and 12 h and processed for analysis as described above at 24 h. The rate of proliferation of bulge cells was calculated on 5-µm-thick transversal frozen sections. Immunolabeling of consecutive sections with K19 with a guinea pig anti-mouse K19 antibody (see below) provided a suitable landmark to identify the bulge area. Percentages of BrdU-positive (cf. above) basal cells and suprabasal cells were measured after 1 month (n=7) and 7 month (n=21) chases. The bilateral Student’s t test was used for statistical analysis.
Immunolabeling for light microscopy
Indirect immunofluorescence was done on 5-µm-thick, acetone-fixed (10 min at –20°C) frozen sections prepared from adult C3H/Hen, FVB, C57BL/6 mice, C57BL/6 K17–/– mice, Fisher rats (Charles River), and tissue-engineered skin as described (5)
. We used rabbit polyclonal antisera raised against K14 (13)
, C-terminal peptide of K14 (14)
, K6 and K17 (15)
, mouse K16 (16)
, mouse K5 (AF138; BabCO, Richmond, CA, USA), human collagen IV (Chemicon, Temecula, CA, USA) and mouse loricrin AF 62 (Covance, Cumberland, VA, USA). Other antibodies used included guinea-pig polyclonals raised against mouse K19 (5)
, K8/18 (American Research Product, Belmont, MA, USA) and human K15 (Progen Biotechnik, Heidelberg, Germany), mouse monoclonal against K14 (Sigma) and a rat monoclonal against laminin (Chemicon). Secondary antibodies used were goat anti-rabbit IgG-IgM or anti-rat IgG (H+L) conjugated with DTAF (fluorescein dichlorotriazine) or Rhodamine (Chemicon) and goat anti-guinea pig IgG (H+L) conjugated with FITC (fluorescein isothiocyanate) (Jackson Immunoresearch Laboratories, West Grove, PA, USA). In some cases, Hoechst stain was applied to visualize nuclei. Following observation, sections were counterstained with hematoxylin-and-eosin. For immunoperoxidase stainings, slides were sequentially incubated with formol (1%, 10 min), methanol (100%, 10 min), NaOH (0.07N, 15 s) and the Vector M.O.M Basic Kit to reduce nonspecific binding. Sections were reacted with mouse monoclonal raised against BrdU and keratin 10 (CK8.60, Sigma), the Ultra horseradish peroxidase detection system (ID Labs Inc, London, ON, Canada), and 3–3'-diaminobenzidine (Sigma). Nuclei were counterstained with Harris’s hematoxylin. A Nikon Eclipse E600 microscope (Nikon, Quebec, QC, Canada) equipped with a SenSys or Coolsnap digital camera for immunofluorescence pictures or color pictures, respectively, was used. Images were processed with the Adobe Photoshop 7.0 software (Adobe Systems, San Jose, CA, USA).
Immunolabeling for transmission electron microscopy
An immunogold labeling protocol was adapted from Blessing et al. (17)
and Franke et al. (18)
. Frozen sections (8 µm thick) of vibrissa follicles from adult (2–3 months old) C57BL/6 K17–/–, wild-type C57BL/6, and FVB mice were incubated for 5 min in 10 mM Tris-HCl (pH 7.2) buffer containing 1% Triton, 1.5 M KCl, and 10 mM EDTA, washed in PBS buffer, and fixed with acetone for 10 min at –20°C. For double-labeling, sections were sequentially incubated with rabbit anti-K14 (13)
, goat anti-rabbit IgG coupled to 10 nm colloidal gold particles (British Biocell International, Cardiff, UK), guinea-pig anti-K19 and goat anti-guinea pig IgG coupled to 5 nm colloidal gold particles (British Biocell International). All steps were separated by PBS-bovine serum albumin rinses. After labeling, the sections were fixed with 2.5% glutaraldehyde for 15 min, washed with cacodylate 0.2 M buffer, postfixed with 2% OsO4 for 30 min, and embedded in LR White. Thin sections stained with lead citrate were observed with a JEOL 1200 EX transmission electron microscope (Jeol Ltd., Tokyo, Japan).
In situ hybridization and terminal deoxynucleotidyl transferase-mediated nick end labeling staining
In situ hybridization was performed on frozen sections prepared from paraformaldehyde-fixed, OCT-embedded whisker pad samples from 1-month-old C57BL/6 mice. Digoxigenin-labeled sense (control) and antisense probes corresponding to 3' end of exon 8 in either mK17 (284 bp), mK16 (208 bp) or mK14 (206 bp) were made according to the MAXIscript protocol (Ambion, Austin, TX, USA) and hybridized to sections. After washes, bound probe was revealed by alkaline phosphatase activity (Bio-Rad, Mississauga, ON, Canada). For mK17, mK16, and mK14, each probe targets a portion of the C-terminal tail domain and most of the 3' noncoding region (16
, 19)
. Terminal deoxynucleotidyl transferase-mediated nick end labeling (TUNEL) staining was performed as described (12)
using 4% paraformaldehyde-fixed paraffin-embedded skin samples.
Analysis of cell proliferation and tissue-engineered skin
To evaluate the colony-forming efficiency of vibrissa bulge cells, 4-week-old Fisher rats (Charles River) were used. According to the classification of Ibrahim and Wright (20)
, the E, F, G, H, Ia-Va vibrissa follicles were dissected under a Nikon Smz-2T binocular microscope. Vibrissa bulge sections were obtained as described (21)
. Dissected bulges (90–120) were put in a 40-µm-pore cell strainer (Falcon, Becton Dickinson, Oakville, ON, Canada) within a 60-mm petri dish containing 10 ml of 1 mg/ml collagenase/dispase (Roche, Laval, QC, Canada), and incubated 30 min at 37°C. The solution was removed and replaced with 0.05% trypsin and 0.01% EDTA. A magnetic bar was deposited in the center of the cell strainer, and bulge fragments were incubated under rotation for 5 min. Afterward, keratinocytes detached from the bulge fragments were harvested along with the medium of the cell strainer. After washing, cells were seeded into 4 or 5 wells (12-well plate, Falcon 353043), containing 25,000 lethally irradiated 3T3 and 1 ml of complete keratinocyte medium as previously described (22)
and incubated in 8% CO2, 100% humidity atmosphere at 37°C. Fresh trypsin was added to the petri dish containing the cell strainer, and bulge fragments were further incubated 5 min. Ten additional periods of trypsin digestion were carried out, and cells were harvested and cultured for each period as described above. To evaluate the effect of the length of time in trypsin, in another set of experiments, cells harvested after 2 and 3 serial trypsinizations were maintained in trypsin for 10 and 15 min longer, respectively. Cells were cultivated for 11 days before staining with 1% rhodanile blue. Colonies were counted as large (i.e., >2 mm diameter), medium (i.e., >1 and <2 mm diameter) or small clones (i.e., <1 mm diameter).
To investigate the tissue engineering potential and epidermal differentiation of vibrissa cells, vibrissa follicles from FVB mice (12 days old) were dissected and fragmented according to Kobayashi et al. (21)
. Ten individual fragments of bulge- or bulb-containing segments or the intermediate region between were explanted onto a tissue-engineered dermal surface of 5 cm2, prepared as described (23)
, and cultured in complete keratinocyte medium containing murine epidermal growth factor (EGF; Sigma). To foster a full epidermal reconstruction, hair explants were placed in submerged cultures for 14 days, at which time the samples were raised at the air-liquid interface for 7 days in EGF-free complete keratinocyte medium. Biopsies were then embedded in OCT. For histological analysis, tissue-engineered skin sections were fixed with Histochoice (Amresco, Solon, OH, USA). Paraffin sections were stained with Masson’s trichrome.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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to compare the potential of various portions of the hair follicle to reconstruct a fully differentiated epidermis in vitro. We explanted dissected bulges, bulbs, and intermediate regions from vibrissa follicles onto tissue-engineered dermis, an excellent substrate for the proliferation of epithelial cells (23)
. Eleven days later, 10/10 bulges, 2/20 bulbs, and 0/10 intermediate regions had generated migrating epithelial tongues (data not shown). Such migration was sustained until confluence (11 days) in bulge explants, but it stopped after 5 days and well prior to reaching confluence in tissues engineered with hair bulb explants.
After 14 days of immerged culture of bulge explants, a 2- to 3-layer-thick epithelium completely covered the dermal surface (5 cm2) (data not shown). After an additional 7 days of culture at the air-liquid interface, a nice epithelium composed of all the strata normally present in interfollicular epidermis, including the granular layer and stratum corneum, had formed (Fig. 1
Aa). This tissue-engineered skin was processed for immunodetection of several markers of epidermal differentiation. Two basal lamina components, collagen IV (Fig. 1Ac
) and laminin (data not shown), were detected at the dermo-epidermal junction as expected. K19 was identified in a small subset of basal keratinocytes (Fig. 1Ad
, open arrowheads), while K15 was expressed in the majority of basal cells (Fig. 1Ae
). K14 (Fig. 1Af
) and K5 (data not shown) were found in the basal and suprabasal living keratinocytes. Bright K17 staining occurred in most basal cells and, in a more sporadic fashion, in lowermost suprabasal cells (Fig. 1Ag
). K10 and loricrin, which are respectively induced at early and late stages of differentiation, occurred as expected from the lower and upper suprabasal layers of the epithelium, respectively (Fig. 1Ah, i
).
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Keratinocytes from the other portions of the vibrissae, such as bulb (Fig. 1Ab
) and intermediate region (data not shown), did not proliferate enough to entirely cover the surface of the tissue-engineered dermis, and failed to form an epidermis. Collectively, these data indicate that cells of the bulge region of mouse vibrissa follicles can proliferate to generate an epithelium exhibiting several of the key properties specific to epidermis and thus are a good cell source for the production of tissue-engineered skin.
Basal and suprabasal cells with distinct histological features within the vibrissa bulge
The bulge area of hair follicles contains multipotent epithelial SCs (see introduction). In vibrissa follicles, the bulge forms a distinct protuberance in the upper third of the hair follicle ORS and comprises multiple cell layers (Fig. 1Ba, d
), making it a convenient setting in which to analyze its cellular constituents. Hematoxylin-and-eosin staining of vibrissa reveals morphological heterogeneity within this region. The histological features of basal (designated Bb for bulge basal cells) and the internal-most suprabasal cells (designated Bs2 for bulge suprabasal 2) are similar to cells making up the ORS in the intermediate region of the vibrissa follicle (Fig. 1Ba, d
). In contrast, the cells located in between these two compartments are markedly more eosinophilic (Fig. 1Ba, d
) and present a denser cytoplasm when viewed by phase contrast microscopy (Fig. 1Be
). These cells, designated Bs1 cells for bulge suprabasal 1, are confined to the bulge area.
Bulge cells possess an extensive proliferation potential: Bb and Bs1 contains LRCs
A distinguishing feature of SCs is their slow-cycling nature (i.e., LRC character). To define the proliferation status of Bb, Bs1, and Bs2 cells, we injected newborn (P1) mice with BrdU using an established protocol, allowing the labeling and localization of LRCs along with the identification of the bulge area by K19 immunolabeling on consecutive sections (24)
. After a chase period of 1 or 7 months, labeled cells were detected in the basal layer of the bulge (Fig. 1Bb, f, g
, arrows), as expected (1
, 5
, 25)
. Moreover, BrdU-labeled cells were also consistently seen in the Bs1 compartment (Fig. 1Bb, f, g
; arrowheads). Neither Bb nor Bs1 cells were actively proliferating since they exhibited no labeling at 24 h following a single BrdU injection (Fig. 1Bc
). Yet, unlike fibroblasts (Fig. 1Bb
, asterisk), these cells have not stopped proliferating altogether, since labeled cells can easily be detected in both the Bb and Bs1 compartments 1 month after repeated BrdU injections to adult mice (Fig. 1Bh
).
To evaluate whether the rate of proliferation of LRCs was similar in Bb and Bs1, the number of LRCs was evaluated after a chase period of 1 or 7 months. Following a 1-month chase, LRCs were present in both Bb and Bs1 compartments (Fig. 1C
). After a 7-month chase period, BrdU-labeled Bs1 cells exhibited a larger decrease than BrdU-labeled Bb ones (Fig. 1C
). These results suggest that the Bb cell population cycle less frequently than the Bs1.
A functional characteristic of SCs is their extensive proliferative potential, although they are quiescent in situ (4
, 21
, 26
, 27)
. Thus, we analyzed the clonogenicity of Bb and Bs1 cells in vitro. Dissected vibrissa bulge areas were subjected to repeated 5-min digestions with trypsin. Following every digestion, cells were harvested and cultured to evaluate their colony-forming efficiency, and fresh trypsin was added to the remaining bulges. Labeling these serially trypsinized bulges for K15, which labels Bb but not Bs1 cells in situ (see below and Fig. 2
Ad, e), showed that most Bb cells were extracted between the first and fourth digestions (Fig. 1Da
, arrows). No K15-bright cells were seen after 6 serial digestions (30 min, Fig. 1Db
), indicating that continued trypsinization yields Bs1 cells at this stage. When testing for growth potential ex vivo, large colonies (i.e., >2 mm diameter) occurred in cultures established from the material harvested after 5, 10, 15 min of serial trypsinization. Large colonies also occurred in cultures seeded from cells harvested after 20 min of serial trypsinization, when very few K15-bright Bb cells persist in the digested bulges, and after 30 min and beyond, when only Bs1 keratinocytes can be harvested. In contrast, only very few small (<1 mm diameter) colonies were found in cultures established from cells obtained after 50 min of serial trypsinization. As a control, the more clonogenic cells (those detached from the bulge after 2 serial trypsinization over 10 min; Fig. 1Df
) were maintained in trypsin for a further 10 min (10+10) before plating. Yet, similar numbers of colonies were obtained (Fig. 1Dg
). A similar outcome was obtained when considering longer trypsinization times (up to 30 min; data not shown), indicating that a prolonged time in trypsin does not significantly reduce colony-forming efficiency. This experiment was conducted twice with similar results. We conclude that both Bb and Bs1 compartments comprise functional SCs.
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Taken together, these findings establish that SCs exhibiting slow-cycling properties occur in the basal, as well as suprabasal compartments within the vibrissa bulge, that both Bb and Bs1 have colony-forming capacity, and that Bs1 suprabasal cells are distinct by virtue of their optically dense and eosinophilic cytoplasm.
Asymmetry of keratin distribution in the SC niche of the vibrissa follicle
To better characterize the SC niche of the vibrissa follicle, we took advantage of the tight differentiation-related regulation of keratin genes within epithelia (9)
. Frozen sections of adult murine vibrissa follicles were immunostained for K19 and K15, which are putative SC markers (5
, 10)
, and for the other main keratins occurring in the ORS, including K5, K6, K14, K16, K17 (28
29
30
31)
. K19 was localized in basal cells of the bulge area (Figs. 1Bi, j
and 2Aa
). A subset of K19-expressing cells (Fig. 2Aa
, arrowheads) reacted with antibodies directed against K8/K18 (Fig. 2Ab
, arrows), indicating that they were Merkel cells (32
, 33)
. Unlike the situation prevailing in epidermis (34
, 35)
, cells within both the Bb and Bs1 compartments were negative for K14 (Fig. 1Bi, j
). K14 immunoreactivity was, however, prominent in the Bs2 compartment of the bulge and in the intermediate region of the ORS (Fig. 1Bi, j
). The use of three antibodies raised against different K14 epitopes yielded similar findings in vibrissa follicles from FVB, C57BL/6 (Fig. 2Cf
) and C3H/Hen mice (data not shown). In situ hybridization for the K14 mRNA yielded identical findings (Fig. 2B
). This distribution did not change significantly as a function of the vibrissa growth cycle (Fig. 2B
; compare a, b with c, d). Remarkably, therefore, the slow-cycling cells contained within the Bb and Bs1 compartments of the bulge do not express K14.
In contrast to K19, immunostaining for K15 gave rise to a signal in all basal keratinocytes of the ORS (Fig. 2Ad
). K15 was also expressed in Bs2 cells in addition to all suprabasal cells in ORS from the intermediate region of the vibrissa follicle. Bs1 cells were distinctly devoid of K15 (Fig. 2Ad
). K16 antigens (Fig. 2Ag
) and mRNA (data not shown) were restricted to the companion layer. In contrast, K17 antigens (Fig. 2Af
) and mRNA (data not shown) occurred throughout the bulge (Bb, Bs1, Bs2 cells alike) and in the remainder of the ORS. The unique character of Bs1 cells, with strong expression of K17 but little if any K14 and K15, was observed in adult rat vibrissae as well (Fig. 2Ac, e, h
).
We also performed immunostainings for conventional K6 isoforms and K5, the type II keratins most frequently coregulated with K17 in vivo (36
37
38
39)
. K5 immunoreactivity occurred in all three compartments of the vibrissa bulge and throughout the remaining ORS (Fig. 2Ca
). On the other hand, K6 protein expression was restricted to the innermost layers of the ORS (Fig. 2Cc
) that also expressed K16 (Fig. 2Ce
). K5 thus represents the likely polymerization partner of K17 in Bs1 cells of the vibrissa bulge. K5/K17 copolymerization has been reported in vitro (40)
and in vivo (36)
. On the basis of type I keratin expression, two distinct populations of LRCs can be defined in the skin SC niche of mouse vibrissa follicles: 1) basal (Bb) keratinocytes expressing K15, K17, and K19 but devoid of K14; and 2) suprabasal (Bs1) cells featuring K17 as their main type I keratin.
Establishing a parallel between keratin protein distribution and filament organization
Transmission electron microscopy in association with immunogold labeling for K19 and K14 was carried out in an attempt to determine whether the differential distribution of keratin proteins has any impact on the ultrastructural organization of keratin filaments. In accordance with immunofluorescence observations, bulge basal (Bb) cells exhibited gold labeling for K19 but not for K14 (Fig. 3
Aa, b), correlating with a loosely organized keratin IF network. In stark contrast, Bs1 cells exhibited a very dense network of heavily bundled filaments that did not stain for either K19 or K14 (Fig. 3Aa', c
). Bs2 cells, which were immunopositive for K14 but not K19, displayed a keratin filament network of intermediate density relative to the Bb and Bs1 cells (Fig. 3Aa'', d
), reminiscent of basal cells in other regions of the ORS (Fig. 3Ae, f
) and epidermis. Thus, the differential distribution of type I keratins is mirrored by striking differences in the density and organization of keratin filaments in the three major subcompartments forming the vibrissa bulge.
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Lack of K17 is associated with hallmarks of epithelial fragility in the vibrissa bulge
Availability of K17-null mice (12)
provides an opportunity to assess the consequences associated with loss of this keratin in vibrissa bulge epithelial cells. K17-null mice lack most of their vibrissae (Fig. 4
Ab). In contrast to the previously reported phenomenon of partial alopecia affecting the pelage coat (12)
, the vibrissa phenotype shows a high and strain-independent penetrance and does not normalize with age (data not shown). Histological analyses showed that vibrissa follicles were frequently altered, including deformation of the hair shaft, defects in the hair fiber (Fig. 4Bb
, open asterisk), and destruction of the ORS, including the bulge. Some K17–/– vibrissa follicles appear histologically normal (Fig. 4Bc
). However, close examination of their bulge area shows that lack of K17 is associated with loss of the Bs1-associated features such as strong eosin affinity (Fig. 4B
, compare a, d with b, c, e, f) and dense cytoplasm under phase contrast microscopy (data not shown). No significant differences between null and wild-type samples were observed in bulge preparations subjected to TUNEL staining (data not shown), suggesting that apoptosis is not a factor in Bs1 cell destruction. Thus, lack of K17 has a different impact on vibrissae and pelage hair follicles with regard to susceptibility to apoptosis (12
, 41)
.
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Additional stainings of vibrissa follicle sections were performed to assess whether loss of K17 causes alterations in keratin markers. K16 was distinctly up-regulated in the innermost (hair-proximal) portion of the Bs1 compartment (Fig. 4Cd
, arrows), instead of being restricted to companion layer (Fig. 2Ce
). The outermost layers of the K17–/– Bs1 compartment were, however, negative for K16 (Fig. 4Cd
, open arrowheads), suggesting that the compensatory induction of K16 taking place in pelage follicles (12)
does not occur in the outermost portion of the Bs1 compartment. Another notable difference was the presence of K14-immunopositive cells in the Bb (basal) compartment of K17–/– preparations (Fig. 4Cb
, arrow). This correlated with the presence of a small number of actively proliferating cells in this layer, as shown by acute BrdU incorporation studies (data not shown). This phenomenon likely reflects an attempt toward tissue repair. The distribution of K19, K15, and K6 was largely the same in K17–/– (Fig. 4Cb-f
) and wild-type vibrissa follicles. K5 staining was reduced, absent, and/or atypical in the Bs1 compartment of K17–/– follicles (Fig. 4Ce
, open arrowheads), supporting the notion that K17 normally acts as a polymerization partner for K5 in this compartment. The distribution of K6 was not significantly altered (Fig. 4Cf
).
The presence of proliferating cells in the Bb compartment prompted us to examine the status of BrdU label-retaining cells in K17–/– vibrissae. We found that LRCs still occurred in the basal layer of the bulge (Bb), as well as in the suprabasal cells corresponding to Bs1 (Fig. 4Bg, h
), indicating that K17 is not required for the presence of LRCs in the Bs1 population. Some K17–/– vibrissae featured an asymmetrically shaped bulge with an apparent decrease in the number of suprabasal layers containing LRCs (data not shown). However, thisdetermination was rendered difficult since the more eosinophilic labeling of Bs1 keratinocytes is absent in K17–/– vibrissae.
Immunoelectron microscopy was performed to further define the alterations in the Bs1 compartment of K17–/– vibrissa bulge. Consecutive tissue sections were processed for K19-immunofluorescence (Fig. 3Ba, b
) and immunogold labeling, providing a suitable landmark to identify the Bb compartment, and for ultrastructural analysis. In contrast to wild type (Fig. 3Bc, e
), the Bs1 suprabasal cells located next to K19-expressing cells in the bulge exhibited a poorly developed IF network in K17–/– vibrissa follicles (Fig. 3Bf, h-j
). Large areas of cytoplasm were completely devoid of filaments in these cells. Desmosomes were retained at sites of cell-cell adhesion (Fig. 3Bi
, des), but Bb and Bs1 suprabasal cells were more flattened and electron-lucent (Fig. 3Bf
). These observations provide an ultrastructural basis for the apparent fragility manifested by K17–/– Bs1 cells, a likely contributor to the vibrissa phenotype in these mice.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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3
4
, 7
, 8)
. Of the three distinct cell populations occurring in the bulge area of mouse and rat vibrissa follicles, two are slow-cycling, a characteristic shared by SCs. One, as expected (5)
, consists of K19-expressing cells lying on a basal lamina (Bb cells), and displaying a wispy keratin IF network when compared to "typical" basal keratinocytes (e.g., epidermis). The other one, unexpectedly, is located suprabasally, i.e., away from the basal lamina (Bs1 cells), and consists of cells featuring an unusually dense keratin IF network correlating with an enriched content in K5/K17. The presence of K17 is required for the distinct cytoarchitecture and the integrity of epithelial cells within the Bs1 compartment, and the presence of vibrissa hair on mouse whisker pads. Both Bb and Bs1 cells are negative for K14, usually a mainstay in the progenitor (basal) compartment of stratified epithelia (35
, 42
, 43)
, and generate holoclones in the context of ex vivo culture. Of potential relevance here, epithelial SCs in mouse embryonic esophagus and trachea are K14-negative (44)
. Interestingly, the induction of K14 in Bb cells at anagen onset during postnatal hair cycling, and after vibrissa plucking, correlates with their active proliferation, and is consistent with a role for K19-expressing SCs in the generation of more differentiated keratinocytes (our unpublished data). Our results further the significance of differential keratin regulation in complex epithelial settings.
Oshima et al. (4)
provided evidence for a sustained migration of SCs, from the bulge to the bulb area (within the ORS), in anagen-stage vibrissa follicles. Only when they have reached their destination, the bulb, do these SCs presumably commit to a particular lineage (e.g., IRS, hair shaft). We report here a faster decrease in the number of LRC in the Bs1, compared to the Bb, compartment over time. This observation could mean that Bs1 cells are more actively involved in the maintenance of hair growth during anagen. Hence, Bs1 cells could give rise to terminally differentiated cells of the hair follicle and epidermis. While studying pelage hair follicles in mice, Blanpain et al. (7)
showed that the basal and suprabasal slow-cycling epithelial cells were each multipotential and capable of long-term self-renewal, two defining characteristics for SCs, despite their distinct molecular signature. The presence of SCs in the suprabasal layer(s) of the bulge, away from the basement membrane, is intriguing given the reported ability to enrich for skin epithelial SCs when selecting for high levels of specific integrins exposed at the cell surface (45
, 46)
.
Specific keratins, namely K19 (5)
and K15 (10)
, are "enriched" in the basal layer of the bulge, correlating with a rather wispy keratin filament network. Low levels of K14 in these cells, as seen in our study (see also ref. 43
) might also be relevant in this regard. This said, Blanpain et al. (7)
did not report on lower K14 levels in the bulge basal keratinocytes of adult mouse pelage follicles, pointing to a potential difference with vibrissa follicles. K19 is now found to be consistently present in a group of basal cells encompassing the bulge (and its slow-cycling cells) in human scalp hair, mouse pelage follicles (5)
, and mouse and rat vibrissa follicles (this study). The physiological significance of these findings is unclear at present.
The second population of slow-cycling keratinocytes in the vibrissa bulge exhibits a dense cytoplasm by phase contrast microscopy and a strong affinity for eosin, both reflecting an unusually prominent network of keratins IFs. Bs1 keratinocytes express K5/K17 as their main keratin pair, i.e., they lack many of the keratins usually found in the ORS, such as K14, K15, and K16 (29
30
31
, 47
, 48)
. The presence of this dense network requires K17 (as does the maintenance of K5 protein), ascertaining the predominant role of the K5/K17 pairing in Bs1 keratinocytes of the bulge. K17 expression occurs in many contexts where epithelial cells are recruited for the purpose of epithelialization (39
, 49
, 50)
. K17 protein itself has been directly implicated in three roles, including mechanical support (12)
, regulation of protein synthesis and cell growth (51)
, and attenuation of TNF-
-induced apoptosis (41)
. Whereas our present findings suggest that compromised structural support, not enhanced apoptosis, accounts for the striking alterations occurring in Bs1 bulge keratinocytes lacking K17, whether other aspects of this keratin’s roles in the cell contribute to this phenotype is unclear. LRCs are still present in the absence of K17 in the Bb and Bs1 compartments of the bulge. It is difficult to conclude on the effect of K17 on SC kinetics since these LRCs likely cycle more frequently in an attempt to regenerate the damaged population. The latter is supported by the emergence of K14 expression, correlating with proliferation, in the Bb compartment.
Moll et al. (34)
previously reported on a high density of K17 staining in keratinocytes surrounding groups of Merkel cells in human epidermis. As the hair bulge is also well endowed in Merkel cells (52)
, the extensively developed IF apparatus promoted by K5-K17 in specialized keratinocytes could thus provide the rigidity needed to optimize the mechanical (and chemical) coupling between hair shaft, Merkel cells, and the sensory neurons surrounding the bulge.
In vitro, mouse (present study), as well as human hair follicle bulges, can produce epidermis in tissue-engineered skin (53
54
55)
. Autologous epidermal equivalents from ORS cells have been used to treat chronic ulcers (56)
. The importance of SCs in the regeneration of the epithelium on a long-term basis is well recognized. Thus, the production of tissue-engineered skin from a region enriched in SCs such as the bulge is likely to present advantages. Additional studies of grafting tissue-engineered skin produced from bulge SCs on athymic mouse will be required to test the hypothesis of long-term survival in vivo before the development of clinical applications. The quality of the epidermis present on tissue-engineered skin and its complete differentiation status suggest that it is a good model for in vitro applications, such as the study of the effects of genetic modifications, natural or transgenic, on epidermal structure and organization in a controlled environment.
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ACKNOWLEDGMENTS |
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Received for publication February 16, 2007. Accepted for publication November 29, 2007.
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  M. A. M. Akinci, H. Turner, M. Taveras, A. Barash, Z. Wang, P. Reinach, and J. M. Wolosin Molecular Profiling of Conjunctival Epithelial Side-Population Stem Cells: Atypical Cell Surface Markers and Sources of a Slow-Cycling Phenotype Invest. Ophthalmol. Vis. Sci., September 1, 2009; 50(9): 4162 - 4172. [Abstract] [Full Text] [PDF]
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 C.-H. Lee and P. A. Coulombe Self-organization of keratin intermediate filaments into cross-linked networks J. Cell Biol., August 10, 2009; 186(3): 409 - 421. [Abstract] [Full Text] [PDF]
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