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. 2015 Oct 1;228(1):153–161. doi: 10.1111/joa.12387

Expression of amelogenin and effects of cyclosporin A in developing hair follicles in rats

Hong‐Il Yoo 1, Gye‐Hyeok Lee 1, Su‐Young Lee 1, Jee‐Hae Kang 1, Jung‐Sun Moon 1, Min‐Seok Kim 1,✉, Sun‐Hun Kim 1,✉
PMCID: PMC4694157  PMID: 26426935

Abstract

Amelogenin, an enamel matrix protein has been considered to be exclusively expressed by ameloblasts during odontogenesis. However, burgeoning evidence indicates that amelogenin is also expressed in non‐mineralizing tissues. Under the hypothesis that amelogenin may be a functional molecule in developing hair follicles which share developmental features with odontogenesis, this study for the first time elucidated the presence and functional changes of amelogenin and its receptors during rat hair follicle development. Amelogenin was specifically localized in the outer epithelial root sheath of hair follicles. Its expression appeared in the deeper portion of hair follicles, i.e. the bulbar and suprabulbar regions rather than the superficial region. Lamp‐1, an amelogenin receptor, was localized in either follicular cells or outer epithelial sheath cells, reflecting functional changes during development. The expression of amelogenin splicing variants increased in a time‐dependent manner during postnatal development of hair follicles. Amelogenin expression was increased by treatment with cyclosporin A, which is an inducer of anagen in the hair follicle, whereas the level of Lamp‐1 and ‐2 was decreased by cyclosporin A treatment. These results suggest that amelogenin may be a functional molecule involved in the development of the hair follicle rather than an inert hair shaft matrix protein.

Keywords: amelogenin, cyclosporin A, development, hair follicle, Lamp‐1

Bullet points

Amelogenin and its receptors are present and modulated in developing hair follicles

Introduction

Amelogenin has been considered to be tissue‐specific and exclusively expressed by (pre)ameloblasts of epithelial origin during tooth development for more than 4 decades. Therefore, the function of amelogenin has been widely investigated for hydroxyapatite crystal nucleation in the developing enamel (He et al. 2008). However, recent reports suggest that this protein is implicated in cementogenesis and periodontal ligament regeneration (Janones et al. 2005; Tanimoto et al. 2012), and wound healing (Romanelli et al. 2008). Knockout of amelogenin induced not only abnormal enamel formation and mineralization (Gibson et al. 2001) but also a progressive deterioration of cementum (Hatakeyama et al. 2003, 2006).

Moreover, burgeoning evidence indicates that this protein is also expressed in cells of non‐mineralizing tissues, including brain glial cells and hematopoietic cells (Deutsch et al. 2006), and in the epiphyseal growth plate (Hatakeyama et al. 2014). Reports suggesting that lysosome‐associated membrane protein 1 (Lamp‐1), a transmembrane protein, serves as a cell surface binding receptor for full length amelogenin and leucine‐rich amelogenin peptide (LRAP), a small amelogenin gene‐spliced product, indicate a possibility that amelogenin may function as a paracrine or autocrine molecule in cell signaling (Tompkins et al. 2006; Zhang et al. 2010). In fact, LRAP has been suggested to have chondrogenic and osteogenic potential (Veis, 2003; Viswanathan et al. 2003).

During the development of the hair follicle, the hair shaft‐producing mini‐organ shares the mechanism of epithelial–mesenchymal reciprocal interactions with tooth development (Botchkarev & Kishimoto, 2003; Pispa & Thesleff, 2003; Rendl et al. 2005; Mou et al. 2006), and furthermore, mesenchymal cells derived from hair follicles have odontogenic potential (Wu et al. 2009). Mounting evidence for amelogenin expression in non‐mineralizing tissues and the binding ability of amelogenin to heparan sulfate, a major proteoglycan, indicates a possibility that amelogenin may function as a potential signaling molecule in the development of the hair follicle (Saito et al. 2004).

Knowledge about the molecules that regulate hair follicle formation and growth is essential for achieving therapeutic goals for hair loss conditions. These goals may include the ability to create new hair follicles, to change the characteristics of existing follicles, and to alter hair growth in existing follicles. The identification of molecules that are capable of inducing the formation of new hair follicles will provide us with potential strategies for treating conditions in which there is a lack of hair. Conversely, inhibiting the activities of molecules important for hair follicle formation and cyclical growth may also ultimately provide us with means for treating hirsutism (Millar, 2002). This study was performed to test the hypothesis that amelogenin may be a functional molecule, and this study for the first time elucidated the presence and modulation of amelogenin and Lamp as its receptor during hair follicle development.

Materials and methods

Animals and administration of cyclosporin A

All procedures were performed in accordance with the ethical standards formulated by the Animal Care and Use Committee of Chonnam National University. Sprague–Dawley rats at prenatal days 15, 17 and 19, postnatal days 0, 1, 3, 6 and 9, and adults were housed in laboratory animal care‐approved facilities. Cyclosporin A (CsA, 10 mg kg−1 of body weight) (Fluka, Buchs, Switzerland) was dissolved in dimethyl sulfoxide (DMSO) and subcutaneously injected into the posterior neck of rat pups at postnatal day 0 for 3, 6 and 9 days every other day. For the control, DMSO alone was administered.

Real time RT‐PCR

RNA was extracted from facial skin and developing second molars using a Trizol Reagent (Invitrogen, Carlsbad, CA, USA). The cDNA was synthesized using the superscript first‐strand synthesis system (Invitrogen) and amplified using the Rotor‐Gene RG‐3000 (Corbett Research, Mortlake, Australia). Amplified cDNA was detected using the SYBR Green PCR Master Mix Reagent kit (Qiagen, Valencia, CA, USA). Ratios of the intensities of the target genes and β‐actin signals were used as a relative measure of the target gene expression. The primer sequences (5′ to 3′) used were as follows: amelogenin (GenBank accession number NM_001271074.1), CAAGGCATCCGCTTAACATGG and TGTTCTGCAAGGGTAGGAGA, generating a product of 70 bp; LAMP‐1 (GenBank accession number U75406), CACGTTCAGCACCTGGACTTG and ACCCTGAAAGCCTGGACTTG, generating a product of 144 bp; LAMP‐2 (GenBank accession number NM_017068.2), CGAAAGGAGAGTATTCTACAGCTCA and TGATGGCGCTTGAGACCAAT, generating a product of 139 bp; β‐actin (Gen‐Bank accession number AB_513429), GATCTGGCACCACACCCTTCT and GGGGTGTTGAAGGTCTCAAA, generating a product of 144 bp.

Immunofluorescence and immunohistochemical staining

Facial skin and parts of the maxilla containing developing upper first molar germs were fixed in a 4% paraformaldehyde solution and embedded in paraffin for preparing tissue sections. A rabbit polyclonal antibody against amelogenin and a mouse monoclonal antibody against Lamp‐1 (Santa Cruz Biotech Inc., Delaware, CA, USA) were used as the primary antibodies. Normal serum was substituted for the primary antibody as the negative control. Immunofluorescence staining was performed using the TSA™ Kit (Invitrogen). Briefly, after blocking endogenous peroxidase, sections were reacted with the primary antibody overnight, and subsequently with the HRP‐conjugated secondary antibody (Cell Signaling Technology, Beverly, MA, USA). The sections were incubated in Alexa Fluor 488® tyramide working solution and then counterstained with propidium iodide for nuclear morphology. Sections were photographed using an LSM confocal microscope (Carl Zeiss, Standort Gȍttingen‐Vertrieb, Germany). Immunohistochemical staining was also performed for the amelogenin detection using Histostain®‐Plus kit (Invitrogen). Sections were reacted with the same primary antibody against amelogenin and subsequently with the biotinylated secondary antibody. They were then bound to HRP‐conjugated streptavidin, followed by red coloring with AEC and counterstaining with methyl green (see Supporting Information Fig. S1).

Western blot

The protein was extracted using the Ready Prep protein extraction kit (Bio‐Rad, Hercules, CA, USA) and transferred to the Protran nitrocellulose membrane (Whatman GmbH, Dassel, Germany). Subsequently, the membrane was reacted with the primary antibodies against amelogenin and Lamp‐1 (Santa Cruz Biotech Inc.), and then with HRP‐conjugated secondary antibody (Cell Signaling Technology). Mouse monoclonal antibody against β‐actin (Sigma‐Aldrich Co., St. Louis, MO, USA) and anti‐mouse secondary antibody (Cell Signaling Technology) were used as a reference. The bound antibodies were reacted with the Lumiglo reagent (Millipore, Billerica, MA, USA). The reactants were visualized and photographed using an LAS 4000 mini loaded with an imagereader las‐4000 software (Fujifilm, Minatoku, Tokyo, Japan).

Results

Histological findings of developing hair follicles

Vibrissae, or whickers, are a large, special type of hair follicle. Developing vibrissae were at a placode or primitive peg stage at prenatal day 15 when epithelial cell proliferation was active (Fig. 1a). At prenatal day 17, they were at an early bulbous peg stage when epithelial cells differentiated into the relatively light‐stained inner epithelial cell mass and dark‐stained outer epithelial cell mass (Fig. 1b). They were further elongated and showed a primitive form of a hair shaft in the most central region at prenatal day 19. Also, small hair follicles were burgeoning from the surface epithelium (Fig. 1c). At postnatal day 1, vibrissae became thicker and the hair shaft with traces of dead cells appeared. Inner epithelial mass showed two definite layers that formed the inner root sheath: the outer single cell layer (Henle's layer) and the inner layer consisting of a few cells (Huxley's layer). The outer cell mass became thicker and formed the outer root sheath composed of several layers of inner cells and a single layer of outer cells. The outer root sheath was continuous with the surface epithelium and it rested on the basal lamina that interfaced with the subjacent fibrous encapsulation. The encapsulation became thicker and was differentiated into two layers: inner and outermost fibrous cell layers (Fig. 1d1,d2). At postnatal day 3, spaces filled with blood cells began to form in the inner fibrous encapsulation (Fig. 1e) and they showed a more definitive outline at postnatal day 6 (Fig. 1f). At postnatal day 9, spaces further formed a single cavity called a blood sinus. Both inner and outer root sheaths became thin and delicate, and showed a fully grown morphology (Fig. 1g). In the adult, a vibrissa showed the same structural layers with a dermal papilla (Fig. 1h).

Figure 1.

Figure 1

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H–E staining of developing rat hair follicles. (a) Vibrissae or whiskers developing from the surface epithelium are at the placode stage at prenatal day 15. Ingrowth of surface epithelium (E) and mesenchymal cell condensation (M) are seen. (b) Many vibrissae are elongated into the mesenchyme and are at the early bulbous peg stage at prenatal day 17. Epithelial cells formed a hair bulb (HB) at the distal end and differentiated into two layers: inner epithelial cell mass (IE) and outer epithelial cell mass (OE). (c) In the magnified inset, a developing vibrissa at prenatal day 19 shows a hair shaft anlage in the center, the inner epithelial cell mass (IE), outer epithelial cell mass (OE) and fibrous encapsulation (FE). (d1) Numerous small hair follicles (arrows) as well as large vibrissa follicles are developing at postnatal day 1. (d2) At postnatal day 1, the inner epithelial cell mass of vibrissae develops into the inner root sheath (IRS), whereas the outer epithelial cell mass becomes thick, forming the outer root sheath (ORS). Basal cells in the ORS rest on the basal lamina that interfaces with two layers of fibrous encapsulation: inner (IFE) and outer (OFE). (e) At postnatal day 3, narrow slits (*) are formed in the inner fibrous encapsulation. (f) At postnatal day 6, definitive vascular spaces develop. Besides, numerous developing small hair follicles are seen. (g) At postnatal day 9, vibrissae develop a blood sinus (BS), outside which many nerve bundles (N) appear. (h) In an adult, a longitudinal section of a hair follicle shows definite layers of IRS and ORS and dermal papilla (DP).

Localization of amelogenin and Lamp‐1

Immunoreactivity for amelogenin in vibrissa follicles at postnatal day 1 was negative in the negative control for which the primary antibody was replaced with normal serum (Fig. 2a). The positive control showed strong reactivity in the developing enamel and ameloblasts from the second molar germs at postnatal day 9 (see Supporting Information Fig. S2). There was hardly any immunoreactivity found for amelogenin at prenatal day 15 (Fig. 2b). The first reactivity was seen in developing vibrissae at the early bulbous peg stage at prenatal day 17 (Fig. 2c). Immunoreactivity was negative in most small hair follicles at prenatal day 19. However, reactivity was seen in the epithelial cell mass in some large developing vibrissae (Fig. 2d). At postnatal day 1, strong reactivity was observed in many vibrissae follicles. The reactivity was specifically localized in the epithelial cells in the outer root sheath of vibrissae in a cross‐section of hair follicles. In a longitudinal section of hair follicles, the reactivity was localized in the deep region of vibrissa follicles, i.e. bulbar and suprabulbar regions, rather than the superficial region, i.e. isthmus and infundibulum. Neither the dermal papilla of mesenchymal origin in the recess of basal bulb nor the surface epithelium showed any reactivity (Fig. 2e1,e2). Immunoreactivity was seen in small hair follicles as well as in large vibrissa follicles at postnatal day 3 (Fig. 2f). Similarly, reactivity was also found in the outer root sheath cells at postnatal day 9 (Fig. 2g). A vibrissa in an adult also showed immunoreactivity in the same layer (Fig. 2h). Immunoreactivity for Lamp‐1, an amelogenin receptor, varied in developing follicles. It was found in either the inner fibrous encapsulation or the outer root sheath cells. Reactivity was also localized in most outer root sheath cells abutting the fibrous encapsulation (Fig. 2i1–i3).

Figure 2.

Figure 2

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Localization of amelogenin (a–h) and Lamp‐1 (i). (a) Postnatal day 1. Immunoreactivity was negative in the negative control. (b) No immunoreactivity can be detected in the developing vibrissa at the placode stage (*) at prenatal day 15. (c) Immunoreactivity is found in bulbous peg‐shaped vibrissae at prenatal day 17. (d) Prenatal day 19. Immunoreactivity was negative in burgeoning vibrissa follicles (B), but relatively large sized follicles (A) show reactivity in the epithelial cell mass of vibrissae. (e1) Postnatal day 1. Reactivity is seen in many cross‐cut vibrissa follicles in the deeper region, whereas it is scarcely found in follicles in the superficial region. (e2) Postnatal day 1. Longitudinally cut vibrissa follicles demonstrate that the deeper region of follicles shows immunoreactivity, whereas neither the basal bulb region (small arrows) abutting dermal papilla (*) nor the superficial region of follicles show any reactivity. A large arrow indicates the surface epithelium. (f) Postnatal day 3. Strong immunoreactivity is seen not only in the outer root sheath cells of vibrissa follicles but also in the small hair follicles (arrows). (g) Postnatal day 9. The reactivity is seen in the outer root sheath cells of hair follicles with a blood sinus. (h) Adult. A cross‐section of a vibrissa shows immunoreactivity at outer root sheath cells (ORS). (i1–i3) Postnatal day 9. Immunoreactivity for Lamp‐1 is seen in either the fibrous encapsulation (i1) or mostly in outer root sheath cells abutting only fibrous encapsulation (i2) or in outer root sheath cells (i3).

Expression of amelogenin and its receptors during hair follicle development

Real time RT‐PCR was performed to compare the level of amelogenin expression during hair follicle development in the face. The mRNA expression was increased up to postnatal day 9 in a time‐dependent manner, followed by maintenance of the expression level in adult rats. However, amelogenin expression in hair follicles was considerably low compared with that in the 2nd molar tooth follicles at postnatal day 9, which were used as a control (Fig. 3a). Amelogenin expression in hair follicles was also confirmed at the protein level, showing many splicing variants which represent the characteristics of amelogenin activation (Fig. 3b).

Figure 3.

Figure 3

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Level of expression of amelognin and its receptors during development. (a) Amplicons of amelogenin were generated from facial skin hair follicles at prenatal day 19, postnatal days 1, 3, 6, and 9, and adult rats. The level of the mRNA expression increased in a time‐dependent manner and it was maintained in adult rats. However, the expression in hair follicles is significantly lower than that in the second molar germs. Real time PCR was conducted three times independently and the results are shown as the mean ± SD. *P < 0.05. (b) Amelogenin in hair follicles was detected as many splicing variants by Western blotting. The second molar tooth germs at postnatal day 9 were used as a positive control and show a higher level of expression and many more splicing variants of amelogenin. (c) Amplicons of Lamp‐1 and ‐2 were generated from facial skin hair follicles at prenatal day 19, postnatal days 1, 3, 6 and 9, and adult rats. The level of the mRNA expression of both receptors was relatively constant during development. Real time RT‐PCR was conducted three times independently and the results are shown as the mean ± SD. *P < 0.05. dpp, days postpartum.

Real time RT‐PCR was also performed to determine the mRNA level of Lamp‐1 and ‐2, receptors of amelogenin, during developmental stages of hair follicles. The level of mRNA expression of both receptors was relatively constant during the early developmental period, except in adult rats. Moreover, the level of mRNA expression in hair follicles was less than that in developing second molar germs used as the positive control (Fig. 3c).

Effects of CsA on expression of amelogenin and its receptors during hair follicle development

CsA has been known to have stimulating effects on hair growth and causes hirsutism. To determine the involvement of amelogenin during CsA‐stimulated hair follicle growth, CsA was subcutaneously administered to rats at postnatal day 1. CsA apparently stimulated hair follicle growth, compared with that in the control rats (Fig. 4a). The amelogenin mRNA expression was significantly increased by CsA treatment for 3 days, but this effect was attenuated at days 6 and 9 after the same treatment (Fig. 4b). The increased expression of amelogenin protein by CsA stimulation was also confirmed by Western blotting (Fig. 4c).

Figure 4.

Figure 4

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Modulation of amelogenin by CsA treatment. CsA (10 mg kg−1 of body weight) was subcutaneously injected into the posterior neck of rat pups at postnatal day 1 for 3, 6 and 9 days every other day. (a) The rat treated with CsA for 3 days show a hairy skin appearance compared with the control rats. (b) The level of the amelogenin mRNA expression in facial skin was significantly increased in the rat treated with CsA for 3 days, followed by attenuation at day 6 and 9. Real time PCR was conducted three times independently and significance was determined at *P < 0.05. (c) The amelogenin protein level in facial skin was determined by Western blotting. The rats that received CsA treatment for 3 and 9 days show a higher level of amelogenin expression than that in the control rat. Dpp, days postpartum.

The Lamp‐1 mRNA expression was significantly decreased, but to a small extent, by CsA treatment for 9 days. The Lamp‐2 mRNA expression was also significantly decreased by CsA treatment (Fig. 5a). The decreased expression of Lamp‐1 protein by CsA stimulation was also confirmed by Western blotting (Fig. 5b).

Figure 5.

Figure 5

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Modulation of amelogenin receptors by CsA treatment. CsA (10 mg kg−1 of body weight) was subcutaneously injected at postnatal day 1 for 3, 6 and 9 days every other day. (a) The Lamp‐1 mRNA expression was significantly decreased, but to a small extent, by CsA treatment for 9 days. The Lamp‐2 mRNA expression was also significantly decreased by CsA treatment. Real time PCR was conducted three times independently and significance was determined at *P < 0.05. (b) The decreased expression of Lamp‐1 protein by the treatment was also confirmed by Western blotting.

Discussion

It is known that amelogenin is expressed not only in enamel‐forming ameloblasts but also in HERS cells, crucial for the differentiation of dental follicle cells, which correspond to hair follicle cells during development. Amelogenin is present to some degree in a solubilized state and presumably has some capacity to diffuse. It may function as a growth factor‐like molecule solubilized in the aqueous microenvironment through autocrine/paracrine pathways, particularly during development and stress‐induced remodeling (Jacques et al. 2014). Full length amelogenin and LRAP, a spliced form of amelogenin, stimulate the WNT signaling pathway in human PDL cells and osteogenesis (Matsuzawa et al. 2009; Warotayanont et al. 2009). EMD® (EMD, Straumann, Switzerland), a predominantly amelogenin‐containing medicament, alters the activity of dental follicle cells (Hakki et al. 2001; Hatakeyama et al. 2006). Clinically, amelogenin has been used to treat bone and periodontal defects and to induce wound healing (Vowden et al. 2007; Miron et al. 2015).

In the present study, vibrissae follicles were mainly used for amelogenin localization, simply because they are larger than the other types of small hair follicles, and thus it is easy to identify the morphology in detail, although amelogenin was detected in small hair follicles. Vibrissa is a special type of hair follicle sealed by a blood sinus which allows the mechanoreceptors to sense extremely small deflections. In the present study, vibrissae were burgeoning at prenatal day 15 and were at the early bulbous peg stage at postnatal day 17 when epithelial cells began to differentiate. Most vibrissae were further elongated into the deep region of the skin and showed morphological features of poorly differentiated epithelial cell masses at prenatal day 19. At postnatal day 1, there was some differentiation of keratinocytes, and definite keratinocyte layers could be identified in the inner and outer epithelial root sheaths. At postnatal day 9, vibrissae showed a mature morphology with blood sinuses as seen in adult rats. The level of amelogenin expression was not constant during the developmental stages, indicating that this molecule may somehow be implicated in the development of hair follicles. Also, Western blotting revealed that amelogenin was detected as many splicing variants frequently found in the developing enamel, implying that amelogenin found in hair follicles has a functional role rather than simply a structural role.

Anatomically, each hair follicle is divided into four distinct regions: bulb, suprabulbar region, isthmus (between the arrector pili muscle attachment and the sebaceous duct entrance), and infundibulum (Sperling, 2007). As the hair follicle bulb reaches the bulbous peg stage during development, several keratinocyte layers develop to form the highly organized mature hair follicle structure (Millar, 2002). Keratinocytes adjacent to the dermal papilla correspond to basal cells elsewhere in the epidermis. Keratinocytes in the hair bulb and dermal papilla cells constitute the cell matrix, and their proliferation, differentiation and upward movement contributes to the growth of hair follicles. Keratinocytes leave the bulb matrix and differentiate into specific cell types of the hair follicle. In the present study, amelogenin was first detected at the early bulbous stage, when cell differentiation began. Amelogenin was localized in keratinocytes in the hair bulb and suprabulbar zone where differentiated cells or cells undergoing differentiation are present. Also, in the cross‐section of the hair follicle, amelogenin was localized in the outer root sheath cells, and not in the inner root sheath cells, which become arranged into concentric layers to form the hair shaft. The outer root sheath is continuous with the epidermis and corresponds to stratum Malpighii. The outer root sheath does not take part in hair formation and its function is currently unknown. These findings at least imply that amelogenin is not an inert hair shaft matrix protein but a functional molecule involved in hair development including epithelial cell differentiation. These findings are also in contrast to amelogenesis, during which inner enamel epithelia or ameloblasts secrete amelogenin, forming an enamel matrix.

The development of vibrissae investigated in this study could be regarded as an active growing anagen stage. Because the outer root sheath maintains its morphology during hair cycles from anagen to catagen, we may expect its expression during whole cycles, although we could not determine the level of expression depending on cyclic stages. Also, the level of amelogenin expression was increased by the CsA treatment, implying that amelogenin may be involved in the functional regulation of the outer root sheath cells. This is evidenced also by the findings that amelogenin expression was not detected in hair germ or placode stage at prenatal day 15, but first in the bulbous peg stage of primitive vibrissae development, which features outer cell mass formation. For confirmation, further studies may be needed using an in vitro system to be able to control hair cycles.

The development and maintenance of the morphologically different layers in hair follicles has been found to be genetically controlled. For example, Notch1 and its ligands Serrate1 and Serrate2 control the differentiation of keratinocytes (Powell et al. 1998; Favier et al. 2000). In mature hair follicles, bone morphogenetic protein (BMP) signaling is crucial for hair shaft differentiation (Kulessa et al. 2000). Noggin, the BMP inhibitor, causes defects in differentiation of the hair shaft cortex and cuticle (Kulessa et al. 2000). Ectopic expression of Wnt3 in the outer root sheath causes hair shaft fragility and elevated expression of several nonintermediate filament proteins in the hair shaft (Millar et al. 1999). In the present study, the level of amelogenin expression varied during hair follicle development. Moreover, the expression was increased by treatment with CsA, which induces hairy skin and is beneficial for alopecia treatment (Açıkgöz et al. 2014), implying that amelogenin may somehow be functionally involved in hair follicle growth.

Little is known about the processes involved in the uptake of extracellular amelogenin by cells. Furthermore, we cannot provide information on how amelogenin functions during the developmental cycle of hair follicles. Lamp‐1, a specific receptor for LRAP was localized not only in the epithelial cells but also in the hair follicle tissues. This different receptor localization may due to slightly different stages of vibrissae development. The presence of Lamp‐1 in hair follicles is consistent with a report that follicle tissue‐derived periodontium‐related cells have the cell surface Lamp‐1 receptor for amelogenin to bind to (Zhang et al. 2010). The present findings tempt us to assume that secreted amelogenin may bind to Lamp‐1 or Lamp‐2 on hair follicle and epithelial cells to induce growth and differentiation (Le et al. 2007; Stahl et al. 2013). However, in contrast to the increased level of amelogenin by the brief CsA treatment, the reduction in its receptors is surprising given that the level of Lamp‐1 or ‐2 was relatively constant during early developmental periods and was not inversely correlated to levels of amelogenin. Thus, it is more intriguing to elucidate how amelogenin works during hair development.

Hair and teeth share common features in development. Both of them develop from adjacent layers of epithelium and mesenchyme via reciprocal interaction between these two tissues (Pispa & Thesleff, 2003; Marja & Mikkola, 2007). Considering the mounting evidence for amelogenin expression in non‐mineralizing soft tissues and the common mechanism of the hair follicle and tooth development, amelogenin can be suggested as a potential signaling molecule involved in the development of the hair follicle. How amelogenin interacts with other functional molecules to induce the growth of hair follicles needs to be studied further. Investigation of the signaling pathways upon amelogenin binding with its receptor protein during hair development is an avenue for future research. Also, the creation of transgenic mice expressing individual amelogenin isoforms, knockout of amelogenin and rescue of the phenotype are needed for its functional assay.

Author contributions

S‐H Kim and M‐S Kim: concept/design, data analysis/interpretation, drafting of the manuscript; H‐I Yoo, G‐H Lee, S‐Y Lee, J‐H Kang, J‐S Moon: acquisition of data, concept/design.

Supporting information

Fig. S1. Immunohistochemical demonstration of amelogenin in developing hair follicles.

Click here for additional data file. (12.3MB, tif)

Fig. S2. Positive control for amelogenin in a developing tooth.

Click here for additional data file. (4.8MB, tif)

Acknowledgements

This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIP) (2011‐0030121) and the National Research Foundation of Korea (NRF) grant funded by the Korean Government (KRF‐2010‐002598). The authors declare no conflicts of interest with respect to the authorship and/or publication of this article.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Fig. S1. Immunohistochemical demonstration of amelogenin in developing hair follicles.

Click here for additional data file. (12.3MB, tif)

Fig. S2. Positive control for amelogenin in a developing tooth.

Click here for additional data file. (4.8MB, tif)

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