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Differentiation of fibroblasts to myofibroblasts and collagen fibrillogenesis are two processes essential for normal cutaneous development and repair, but their misregulation also underlies skin-associated fibrosis. Periostin is a matricellular protein normally expressed in adult skin, but its role in skin organogenesis, incisional wound healing and skin pathology has yet to be investigated in any depth. Using C57/BL6 mouse skin as model, we first investigated periostin protein and mRNA spatiotemporal expression and distribution during development and after incisional wounding. Secondarily we assessed whether periostin is expressed in human skin pathologies, including keloid and hypertrophic scars, psoriasis and atopic dermatitis. During development, periostin is expressed in the dermis, basement membrane and hair follicles from embryonic through neonatal stages and in the dermis and hair follicle only in adult. In situ hybridization demonstrated that dermal fibroblasts and basal keratinocytes express periostin mRNA. After incisional wounding, periostin becomes re-expressed in the basement membrane within the dermal-epidermal junction at the wound edge re-establishing the embryonic deposition pattern present in the adult. Analysis of periostin expression in human pathologies demonstrated that it is over-expressed in keloid and hypertrophic scars, atopic dermatitis, but is largely absent from sites of inflammation and inflammatory conditions such as psoriasis. Furthermore, in vitro we demonstrated that periostin is a transforming growth factor beta 1 inducible gene in human dermal fibroblasts. We conclude that periostin is an important ECM component during development, in wound healing and is strongly associated with pathological skin remodeling.
Summary: Periostin is a fibrogenic protein that mediates fibroblast differentiation and extracellular matrix synthesis. Here, we show that periostin is dynamically and temporally expressed during skin development, is induced by TGF-β1 in vitro and is significantly upregulated during wound repair as well as cutaneous pathologies.
Originally identified as an 811-amino acid protein secreted by osteoblasts (Takeshita et al. 1993), periostin is an extracellular matrix (ECM) protein containing four domains with structural homology with the insect protein fasciclin-1 (Conway and Molkentin 2008). Periostin was recently classed as a matricellular protein (Norris et al. 2008a) (modulator of cell-matrix interactions and cell function (Bornstein and Sage 2002)) as a result of an explosion of research in the past few years that has shown that periostin is prominently expressed during ECM remodeling, including in heart after myocardial infarction (Kuhn et al. 2007; Shimazaki et al. 2008), asthma-associated sub-epithelial fibrosis in lungs (Takayama et al. 2006), and pulmonary vascular remodeling (Chen et al. 2006). Moreover, periostin is known to be a key regulator during cardiac development which is particularly evident in the atrioventricular valve where a lack of periostin inhibits differentiation of the cushion mesenchyme into myofibroblastic-valve tissue (Butcher et al. 2007; Lie-Venema et al. 2008; Norris et al. 2008b). Initial assessment of adult skin in periostin−/−mice highlighted significant abnormalities in collagen fibrillogenesis which manifest in increased stiffness and decreased elasticity in comparison with wild-type mice (Norris et al. 2007).Therefore, during development, it appears that periostin could be required to mediate differentiation of fibroblasts to myofibroblasts as well as collagen synthesis and assembly.
Although critically important during development, matricellular proteins are typically restricted to tissue remodeling and wound repair in the healthy adult (Hamilton 2008). Unlike many other members of the matricellular protein family, periostin is normally expressed in adult tissues, including skin where it localizes to dermal fibroblasts, keratinocytes and the basal lamina (Jackson-Boeters et al. 2009). Interestingly, we have previously shown that periostin is only present in the extracellular matrix under tissue remodeling conditions such as those associated with pathological insult (nevus) (Jackson-Boeters et al. 2009). To further investigate periostin expression during tissue remodeling, using a full-thickness excisional wound healing model in C57/BL6 mice, we observed periostin in the granulation tissue at 3 days, with protein levels peaking at 7 days and returning to basal levels at 28 days (Jackson-Boeters et al. 2009). Interestingly, maximal periostin expression was associated with the presence of myofibroblasts (Jackson-Boeters et al. 2009), providing further evidence that periostin likely mediates fibroblast to myofibroblast differentiation.
Having shown association of periostin with normal skin, wound repair and nevus (Jackson-Boeters et al. 2009), we hypothesize that periostin is an important mediator of skin development, healing and remodeling. The aim of this study was to examine the spatiotemporal expression patterns of periostin in murine skin during development and incisional wound healing and whether or not persistence of periostin is associated with human cutaneous pathologies. Here we demonstrate a dynamic pattern of expression in the dermis, hair follicles as well as deposition along the basement membrane (BM) within the dermal-epidermal junction (DEJ) during skin organogenesis. Moreover, the abundance of periostin in fibrotic scars and its relative absence in pre-lesional wounds suggest that it may be a key regulator of ECM synthesis and deposition in skin.
Periostin mice (Snider et al. 2008) were maintained under specific-pathogen-free conditions in individual cages with a 12 h light/dark cycle. The animal use protocols were approved by the Institutional Animal Care and Use Committee’s at IUPUI and the University Council on Animal Care at the University of Western Ontario. Serological analyses were performed on the mice prior to experiments to test for the presence of blood borne pathogens or infection. Embryos were collected from timed matings at E13.5, 15.5, 17.5, P2, P9, P19 and P60 and processed for routine paraffin or cryosection sections (10 µm). Skin from postnatal animals was spread on X-ray film (to maintain flat orientation) prior to 4% paraformaldehyde fixation.
Mice were anesthetized with 1.2% Avertin (125 mg/kg) and the back shaved and sterilized using a 70% alcohol swab. A full thickness incision wound (1 cm long) was cut along back skin below the shoulder blades to prevent self-licking. Ointment (100 mg pure white petrolatum jelly; Vaseline) was applied to the wound and changed every 2 days. The mice were caged individually with regular light/dark cycles. The animals were sacrificed at specified post-surgical time points and skin samples were collected as above for histological analysis.
Formalin fixed biopsy specimens were provided by Dr. Jeff Travers (Department of Dermatology, IUPUI). Four samples from patients (mean age, 33.0 years) with severe atopic dermatitis, five samples from patients of psoriasis (mean age, 39.7 years), three samples from patients with keloid scars (mean age, 28 years), 4 samples from patients with hypertrophic scars (mean age, 32.6 years), 2 samples from patients with chronic dermal inflammation (mean age, 64.5 years) and five samples from healthy individuals (mean age, 48.2 years) were included in this study. The fixed human skin samples were processed for routine paraffin section.
Masson’s trichrome staining was performed as previously described to visualize collagen (blue) and ECM (red) deposition(Liu et al. 2008).
Periostin and Ki67 were detected on paraffin sections using the ABC kit (Vectorstain) following the manufacturer’s instructions. The antibodies were diluted 1:3000 for rabbit anti-periostin (Kruzynska-Frejtag et al. 2004), and 1:500 for mouse anti-Ki67 (DB). Signals were revealed by using DAB and hydrogen peroxide as the chromogen. Sections were counterstained with methyl green. As a negative control for periostin labeling specificity, corresponding tissues from age-matched periostin null mice was processed in parallel. The negative control for Ki67 was set by using normal rabbit or mouse serum respectively at 1:500 dilutions. In situ hybridization for periostin mRNA was performed on paraffin sections using S35-labeled anti-sense riboprobes (Lindsley et al. 2007) and the specificity was controlled by using corresponding sense probe.
Human dermal fibroblasts were isolated using an explant technique as previously described (Chen et al. 2008). Cells were cultured in high glucose DMEM (Invitrogen, Burlington, Ontario). All media was supplemented with 10% FBS and 1% antibiotic/antimycotic solution. Cells were plated onto a six well plate at a density of 60,000 cells/well and were allowed to grow for 24 h at 37°C. Cells were then serum-starved for 24 h, pre-treated Transforming growth factor β1 (TGF- β) (R and D Systems, 4 ng/ml) for 6 h (for mRNA analysis) or 6 and 24 h (for protein analysis).
Total RNA was isolated using Trizol reagent (Invitrogen). Total RNA (25 ng) was amplified using the TaqMan One Step RT-PCR Master Mix (4309169; Applied Biosystems Inc., Streetsville, ON, Canada). Reverse transcription and quantitative real-time PCR reactions were performed using the Prism 7900 HT Sequence Detector (Applied Biosystems Inc.). Samples were incubated at 48°C for 30 mins to make cDNA templates. The resulting cDNA was amplified for 40 cycles. Cycles alternated between 95°C for 15 s and 60°C for 1 min. Results were analysed using SDS v2.1 software (Applied Biosystems Inc.). The ΔΔCt method was used to calculate gene expression levels relative to GAPDH and normalized to control cells. Data were log-transformed prior to analysis by one-way analysis of variance and Tukey’s post-hoc test, using Graphpad Software v. 4 (Graphpad Software, La Jolla, CA, USA).
We examined periostin protein deposition in wild-type skin during embryonic development and at several postnatal stages. At E13.5 and E15.5, periostin was faintly detectable along the BM (Fig. 1a, b) at the dermal-epidermal junction (DEJ) and in the lower dermis. At E17.5, the deposition was more uniform and intense in both the DEJ and dermis (Fig. 1c) compared to earlier stages. The robust deposition along DEJ persisted to postnatal day 19 (Fig 1d, e, f). From P9 onwards, the ECM surrounding hair follicles labeled positively for periostin (Fig. 1e, f, g). In adult mice (P60 and older), hair follicles were intensively labeled, but the DEJ labeling and deep dermal staining was significantly reduced (Fig. 1g).
To assess what cell populations produce periostin, we utilized in situ hybridization. Skin was examined from wild-type mice at E17.5, when periostin protein is present in the dermis and DEJ. Basal keratinocytes above the DEJ contained mRNA for periostin (Fig. 2a), as did the dermal fibroblasts/mesenchymal cells within the upper and lower dermis. Using immunohistochemistry, periostin protein deposition closely correlated to the same area (Fig. 2b). However, in hair follicles, cells contained mRNA for periostin (Fig. 2c), but the protein was not present showing that these cells were not actively secreting periostin protein into the ECM (Fig. 2d).
To assess patterns of periostin expression in wound repair, we induced the wound healing process using the established incisional wound model. At day 5, keratinocytes labeled positively for periostin protein, but not the granulation tissue in the wound (Fig 3a, b). Periostin protein was present in the DEJ in areas adjacent to the wound (Fig. 3c). In areas away from the wound site, periostin labeling is decreased suggesting that periostin is upregulated in the DEJ at the wound edges (Fig. 3d). As cell proliferation is a necessary event in re-epithelialization, we examined the expression of the cell proliferation marker Ki67 and periostin on adjacent sections. Significantly, the epithelium within the wound was positive for Ki67 labeling (Fig 3e), as was the tissue at the edge of the wound (Fig. 3f).
As periostin is implicated in fibrosis, we investigated whether periostin levels were altered in several types of skin lesions. Using immunohistochemistry, we examined spatial deposition of periostin in healthy skin, keloid scars, hypertrophic scars, chronically inflamed skin, atopic dermatitis and psoriasis lesions. Periostin protein levels were highest in keloid and hypertrophic scars, but were significantly reduced in chronically inflamed tissue (Fig. 4). In hypertrophic scars, periostin immunoreactivity was associated with large collagen bundles (shown by Masson’s trichrome), but in hypertrophic scars, periostin was present mainly surrounding cells (Fig. 4). The lack of periostin in chronically inflamed tissue correlated with an increase in inflammatory cell infiltration (Fig. 4) as we have previously reported in murine wound repair(Jackson-Boeters et al. 2009). In healthy and psoriatic skin periostin protein was predominantly deposited along the DEJ, but in skin from atopic dermatitis, periostin was detected throughout the entire dermis (Fig. 5).
Fibrosis is often associated with elevated TGF-β levels and periostin is known to be regulated by TGF-β in osteoblasts (Horiuchi et al. 1999), periodontal ligament fibroblasts (Rios et al. 2008) and cardiac valve precursor cells (Norris et al. 2009). We therefore assessed whether periostin is regulated by TGF-β in human dermal fibroblasts. Human dermal fibroblasts were cultured for 6 and 24 h in the presence of 10 µM TGF-β. Periostin mRNA levels were elevated 2-fold 6 h post stimulation (Fig. 6a), and protein levels were elevated at 6 and 24 h post stimulation (Fig. (Fig.66b).
Skin, the largest organ of the body, provides a physical and immunological barrier against pathogens, regulates body temperature and water loss, and mediates endocrine and sensory functions (Boulais and Misery 2008; Loewenthal 1963; Loewenthal 1964). The composition of the ECM is an important determinant of skin homeostasis, and alterations of the ECM constituents can result in abnormal tissue function (Li et al. 2007; Shibusawa et al. 2008). We have recently confirmed that periostin is expressed in human and murine skin (Jackson-Boeters et al. 2009), and periostin knockout mice display a reduction in dermal thickness (Norris et al. 2007). As these findings suggest that periostin is required for normal skin development and homeostasis, we first investigated the spatiotemporal expression of periostin during development and after incisional wound healing.
Using immunohistochemistry, we identified that periostin localizes to the dermis, basement membrane and hair follicles during embryonic development, but is downregulated in all structures except hair follicles in postnatal development. This suggests that periostin may mediate initial development of the dermis and basement membrane. As highlighted, periostin has already been implicated in fibroblast to myofibroblast differentiation in cardiac development (Norris et al. 2009), but our data also implicates periostin in basement membrane and basal keratinocyte development. Periostin has been heavily implicated in epithelial to mesenchymal transition (Kanno et al. 2008; Ruan et al. 2009; Yan and Shao 2006), but our studies suggests that it may influence epithelial behaviour without inducing fibroblast differentiation. It will be of great interest to assess the influence of periostin on keratinocyte behaviour in vitro, as well as assessing where periostin localizes within the basement membrane in vivo.
As periostin is heavily implicated in ECM remodeling, we investigated the expression of periostin after incisional wounding. Periostin was re-expressed in the BM at the wound edges, which correlated with increased keratinocyte proliferation. As keratinocyte migration is required for re-epithelialization of the wound, this suggests that periostin may trigger this process. Alternatively, as matricellular proteins are known to stimulate an intermediate state of cell adhesion associated with increased migration, periostin may act in this capacity stimulating keratinocyte migration into the wound area. Previous studies have shown that periostin is a potent inducer of cell migration in C3H10T1/2 cells (Lindner et al. 2005), epithelial ovarian carcinoma (Gillan et al. 2002), and vascular smooth muscle cells (Li et al. 2009). Interestingly, periostin is not upregulated in the dermis after incisional wounding, particularly when compared to excisional wounding where it is abundant at 3 and 7 days (Jackson-Boeters et al. 2009). This suggests that periostin expression may not be required unless severe trauma occurs to the dermis. Future studies will focus on whether the incisional and excisional wound repair process is altered in periostin null mice.
During wound repair and certain pathologies, changes occur in the composition of the matricellular ECM to provide cell–matrix signals and misregulation of the changes can result in development of various pathologies (Midwood, Valenick et al. 2004; Berk, Fujiwara et al. 2007; Darby and Hewitson 2007). Periostin is prominently expressed during pathological ECM remodeling, including in heart tissue after myocardial infarction (Kuhn et al. 2007; Shimazaki et al. 2008), asthma-associated sub-epithelial fibrosis in lungs (Takayama et al. 2006), and pulmonary vascular remodeling (Chen et al. 2006). We have now confirmed that periostin protein is abundant in fibrotic scars, but not in chronically inflammed skin. Interestingly we have previously shown in excisional wounds in mice that periostin expression is associated with myofibroblasts, but not CD68 positive inflammatory cells (Jackson-Boeters et al. 2009). It is of particular interest that both keloid and hypertrophic scars contain abundant periostin because the structure of the scars is very different: thick collagen bundles are more abundant in hypertrophic scars often forming acellular nodes in the deep dermis (Figs. 4, ,5)5) In contrast, the centre of the keloid lesion has relatively few cells. It will be of significant interest to assess the cellular source of periostin in each scar type using ISH.
We have also validated previous microarray data that reported periostin expression levels (along with 17 other genes including the matricellular protein tenascin-C) are all elevated in atopic dermatitis relative to psoriasis (Nomura et al. 2003). Though acquired periostin deposition has been noted in bronchial asthma (Takayama et al. 2006), the elevated expression levels and altered presentation patterns of periostin within atopic dermatitis are not sufficient to suggest a causal relationship between elevated levels of periostin and atopic dermatitis. Given that we have shown that periostin is a TGF-β1-responsive gene in dermal fibroblasts (Fig. 6), the elevation of periostin expression in atopic dermatitis may simply reflect disturbance of TGF-β signaling that is thought to suppress inflammatory responses in the skin (Sumiyoshi et al. 2002). Nevertheless, periostin expression may serve as a valuable diagnosis marker to differentiate atopic dermatitis from other skin lesions.
In summary, our study has established periostin as a novel dynamic ECM component associated with skin morphogenesis, homeostasis and wound healing. Our data suggests that periostin may exert effects over cells in the epidermis, dermis as well as within the hair follicles at different stages of development. The re-establishment of periostin expression in the basement membrane during wounding further suggests that periostin is an important regulator of keratinocyte biology. While the exact functions of periostin in development of mouse skin and the mechanism governing the dynamic protein deposition remains elusive, the periostin null mice will be of great value to validate the postulated functions of periostin during organogenesis, wound healing and development.
We thank Prof. Jeffery B. Travers (Departments of Dermatology and Pediatrics, IUPUI) for providing the fixed skin biopsy specimens and Linda Jackson-Boeters (Department of Pathology, UWO) for advice on histological analysis. This work was supported, in part, by the National Institutes of Health (S.J.C), the IU Department of Pediatrics (Cardiology) and Riley Children’s Foundation (H-M.Z), Natural Sciences and Engineering Research Council (C.E) and the Canadian Institutes of Health Research IMHA operating grants (D.W.H).
Conflict of interest None
Douglas W. Hamilton, Email: ac.owu.hciluhcs@notlimaH.salguoD.
Simon J. Conway, Email: ude.iupui@yawnocis.