Activin is a member of the transforming growth factor β (TGF-β) family, which plays a crucial role in skin morphogenesis and wound healing. To gain insight into the underlying mechanisms of action, we searched for activin-regulated genes in cultured keratinocytes. One of the identified target genes encodes Id1, a negative regulator of helix–loop–helix transcription factors. We show that Id1, Id2, and Id3 are strongly downregulated by activin in keratinocytes in vitro and in vivo. To determine the role of Id1 in keratinocyte biology, we generated stable HaCaT keratinocyte cell lines overexpressing this protein. Our results revealed that enhanced levels of Id1 do not affect proliferation of keratinocytes in monoculture under exponential culture conditions or in response to activin or TGF-β1. However, in three-dimensional organotypic cultures, Id1-overexpressing HaCaT cells formed a hyperthickened and disorganized epithelium that was characterized by enhanced keratinocyte proliferation, abnormal differentiation, and an increased rate of apoptosis. These results identify an important function of Id1 in the regulation of epidermal homeostasis.
Activins are members of the transforming growth factor β (TGF-β) superfamily that exist as homo- or heterodimers consisting of βA and βB chains. The major forms of activin are activin A (βAβA), activin B (βBβB), and activin AB (βAβB) (Massague, 1990). Activin activities are mediated by heteromeric receptor complexes, consisting of type I (ActRIA/Alk2, ActRIB/Alk4, and Alk7) and type II (ActRII and ActRIIB) receptors, which are all serine/threonine kinases (Mathews and Vale, 1993; Tsuchida et al., 2004). In addition, activins bind to the soluble glycoprotein follistatin that inhibits activin action (Nakamura et al., 1990).
Recent studies provided evidence for a role of activin in skin morphogenesis, repair, and disease. Thus, transgenic mice overexpressing the activin βA chain in the epidermis were characterized by dermal fibrosis and by a hyperthickened epidermis (Munz et al., 1999). After skin injury, activin βA and βB mRNAs and activin A protein levels increase significantly (Hübner et al., 1996; Cruise et al., 2004), suggesting an important role of activin in wound repair. Indeed, transgenic mice overexpressing the activin antagonist follistatin in the epidermis are characterized by strongly delayed wound healing (Wankell et al., 2001a). By contrast, mice overexpressing activin in the epidermis showed enhanced granulation tissue formation and re-epithelialization after skin injury (Munz et al., 1999). These findings demonstrate that aberrant expression and activity of activin are associated with abnormal skin development and impaired wound healing and suggest that activin regulates migration, proliferation, and differentiation of many cell types in the skin, including keratinocytes. This was confirmed by our recent results, which demonstrated that activin receptor signaling in keratinocytes is important for hair follicle morphogenesis and wound re-epithelialization (Bamberger et al., 2005). In addition, a role of activin in the regulation of keratinocyte proliferation and differentiation was found in in vitro studies with keratinocyte monocultures, where a weak inhibitory effect of activin on keratinocyte proliferation and induction of keratinocyte differentiation by this factor was demonstrated (Shimizu et al., 1998; Seishima et al., 1999). In addition, activin was shown to stimulate keratinocyte migration (Zhang et al., 2005).
To gain insight into the mechanisms of activin action in the skin, we searched for genes that are regulated by activin A in cultured keratinocytes. One of the activin target genes that we identified encodes Id1. Id proteins (Id1–Id4) are negative regulators of basic helix–loop–helix (bHLH) transcription factors. They promote cell growth, inhibit differentiation, and play critical roles in development and cancer (Lasorella et al., 2001). Expression of Id proteins is increased in a large variety of human tumors, including cutaneous squamous cell carcinoma and basal cell carcinoma (Langlands et al., 2000; Chaturvedi et al., 2003; Sikder et al., 2003). In addition, enhanced levels of Id1 were detected in biopsies from lesioned psoriatic skin, suggesting a regulatory role of this protein family in keratinocyte proliferation and differentiation (Bjorntorp et al., 2003).
Id genes are known targets of TGF-β and BMP signaling, and they are regulated in a cell-type-specific manner by these factors (Korchynskyi and ten Dijke, 2002; Kurisaki et al., 2003; Kowanetz et al., 2004). However, their physiological roles in the TGF-β/BMP signaling pathways are still poorly understood. Here, we show that activin A suppresses the expression of Id1, Id2, and Id3 in keratinocytes at the mRNA and protein level in vitro and in vivo, and we provide evidence for an important regulatory function of Id1 in skin homeostasis.
Regulation of Id1, Id2, and Id3 expression in HaCaT cells by activin A and TGF-β1
To identify activin-regulated genes in keratinocytes, we hybridized a cDNA array with radioactively labeled cDNA from quiescent HaCaT cells and from cells that had been treated for 5 or 8 h with activin A. Among the 588 genes represented on this filter, only three appeared to be regulated by activin A. One of them encodes Id1, a negative regulator of bHLH transcription factors (Benezra et al., 1990). To verify this regulation and to determine, if other Id family members are also regulated by activin, we treated quiescent human HaCaT keratinocytes with activin A for different time periods. Since TGF-β is known to regulate id genes in different cell types (Ling et al., 2002; Kang et al., 2003; Kurisaki et al., 2003), we used TGF-β1 as a positive control. As shown by RNase protection assay (RPA) (Figure 1a), id1, id2, and id3 mRNA levels were downregulated within 5–8 h of treatment with either activin A or TGF-β1. The time course and extent of downregulation were different for the two TGF-β family members. In the case of id1, the TGF-β1- and activin-induced downregulation was preceded by a transient upregulation between 1 and 3 h after addition of the factor (Figure 1a and data not shown).
To confirm the activin-mediated Id1 regulation at the protein level, total cell lysates from activin A- or TGF-β1-treated cells were subjected to Western blot analysis. Consistent with the results obtained at the RNA level, an initial upregulation of Id1 expression was observed within the first hour after activin- or TGF-β1 treatment (data not shown), followed by a strong downregulation of this protein within 3–8 h (Figure 1b). To verify that this effect is not a starvation artefact, lysates were also prepared from cells, which had been incubated in serum-free medium in the absence of activin. The latter displayed high levels of Id1 protein, and no downregulation was observed upon starvation (Figure 1b). This result was reproduced in several independent experiments (data not shown).
Since the regulation of Id1 expression by TGF-β occurs in a cell-type-specific manner (Chambers et al., 2003; Kurisaki et al., 2003), we tested if this is also the case for activin. Therefore, we treated primary human dermal fibroblasts with either activin A or TGF-β1 for different time periods and subjected the cell lysates to Western blotting. In contrast to keratinocytes, we observed a strong upregulation of Id1 protein expression in dermal fibroblasts after 3 h of activin or TGF-β1 treatment (Figure 1c). Expression subsequently declined, but was still higher compared to controls within 5–8 h after addition of either activin or TGF-β1. Thus, activin-mediated Id1 expression indeed occurs in a cell-type-specific manner. This result was reproduced in an independent experiment (data not shown).
Reduced expression of id1, id2, and id3 genes in tail skin of transgenic mice overexpressing activin in the epidermis
To determine if the three Id family members are also regulated by activin in vivo, we performed RPAs with RNAs from tail skin of transgenic mice that overexpress the activin βA chain under the control of the K14 promoter in the epidermis (Munz et al., 1999). RNAs from wild-type littermates were used as a control. Tail skin from four mice per genotype was pooled for each experiment. Indeed, we found reduced levels of id1, id2, and id3 mRNAs in tail skin of transgenic mice compared to control mice. Figure 1d shows the result of one representative experiment, which was reproduced at least once with independent RNA samples from additional tail biopsies.
Activin-mediated downregulation was also observed on the protein level for Id3 by immunofluorescence staining of tail skin sections from activin-overexpressing and wild-type animals. In normal tail skin, we found strong signals for Id3 in keratinocytes of the epidermis and the hair follicles (Figure 1e, left panel). In tail skin from activin transgenic mice, this keratinocyte staining was almost completely absent, whereas the weak signal in the underlying dermis remained (middle panel). This suggests that activin also suppresses Id3 expression in keratinocytes in vivo. The specificity of the staining was confirmed by preincubation of the antibody with the immunization peptide (right panel).
Taken together, these results demonstrate that overexpression of activin is associated with downregulation of Id1 levels in vivo, indicating that high levels of activin can indeed regulate Id1 expression not only in vitro but also in vivo.
Expression of Id1, Id2, and Id3 in healing skin wounds
We next addressed the question if endogenous activin can also regulate id expression in vivo. For this purpose, we analysed the expression of these transcriptional regulators during the healing process of full-thickness excisional mouse wounds (Figures 2 and 3), since activin is highly expressed between day 1 and day 7 of wound repair (Hübner et al., 1996). The time points that we chose for the expression analysis are characteristic for the three different phases of repair. In the early inflammatory phase, neutrophils and macrophages become activated and enter the wound area (days 1–3). The second phase is characterized by the formation of new tissue and comprises granulation tissue formation and re-epithelialization (days 3–7). The third phase is the phase of tissue remodeling (starting around day 7) (Martin, 1997; Singer and Clark, 1999). Overviews of wounds in phases 2 and 3 are schematically depicted in Figure 3, top panel. As shown in Figure 2, id2 and id3 mRNA levels were slightly downregulated between day 3 and day 7 after injury. Id1 mRNA was regulated differently compared to id2 and id3, since its level of expression was low in unwounded skin and increased transiently early after wounding, starting within 1–3 h after injury (data not shown). Thereafter, it returned to basal levels (Figure 2). At 14 days after injury, wounds were completely re-epithelialized, but still contained a cell-rich granulation tissue. At this stage of repair, id2 and id3 mRNA levels were as high as in unwounded skin, whereas id1 mRNA levels increased again.
We subsequently localized Id proteins in normal and wounded mouse skin by immunofluorescence. Consistent with data obtained with human skin (Langlands et al., 2000; Bjorntorp et al., 2003; Chaturvedi et al., 2003), we found the three Id proteins predominantly in basal keratinocytes of tail skin from BALB/c mice (data not shown). In 5d and 14d skin wounds, Id proteins were expressed in cells of the granulation tissue as well as in the hyperproliferative epithelium, where nuclear as well as cytoplasmic staining was observed (Figure 3). Expression was strongest in basal and lower suprabasal cells. Interestingly, the Id expression pattern showed an inverse relationship to the one of activin that is predominantly found in the upper, more differentiated cells of the wound epidermis (Hübner et al., 1996; Wankell et al., 2001b; SW, unpublished data).
Generation of Id1-overexpressing HaCaT cells
To determine the importance of the activin-mediated downregulation of Id1 for keratinocyte proliferation and differentiation, we generated stable HaCaT cell lines, which overexpress Id1 with a carboxyterminal epitope of the influenza virus hemagglutinin (HA), designated Id1-HA. Id1 was chosen for further analysis because it is highly expressed in wounded and psoriatic skin (see Figure 3 and Bjorntorp et al., 2003). Western blotting was performed with protein lysates from Id1-HA-transfected and vector-transfected (neo) cell clones. In all clones, the levels of Id1-HA were similar to or lower than the levels of endogenous Id1. The strongest expression of the recombinant protein was seen in clone #1, which showed equal levels of endogenous and overexpressed protein under exponential growth conditions (Figure 4a, left panel). However, the expression levels of this clone declined upon culturing and freezing/thawing, suggesting that the high-expressing cells are gradually lost (data not shown). In the other clones, the levels of endogenous Id1 protein exceeded the levels of recombinant Id1-HA. These ratios between recombinant and endogenous Id1 were verified at the RNA level by RPA using a probe that can distinguish between mRNAs encoding endogenous id1 or recombinant id1-HA mRNAs (Figure 4a, right panel). In some clones, we found enhanced levels of endogenous Id1 protein in Id1-HA-expressing clones, but this was not consistently observed (Figure 4a left panel and data not shown).
We next determined if the overexpression of Id1-HA is more pronounced under conditions where the expression of endogenous Id1 is downregulated, for example, upon treatment with TGF-β1. As expected, TGF-β1 treatment resulted in a downregulation of endogenous Id1. By contrast, expression of the CMV promoter-driven id1-HA transgene was suppressed upon starvation (0 and 8 h controls) and subsequently upregulated in response to TGF-β1 (Figure 4b for clone 1). Thus, the recombinant protein was indeed strongly overexpressed in TGF-β1-treated keratinocytes.
Since Id expression is downregulated during differentiation of other cell types, we determined if this is also the case for HaCaT keratinocytes. For this purpose, cells were grown to confluence and subsequently maintained in the same medium for several days. Under these conditions, partial differentiation of HaCaT keratinocytes is achieved (Ryle et al., 1989). In Western blot experiments, we indeed observed a differentiation-induced downregulation of Id1 (Figure 4c) that is consistent with data obtained with primary keratinocytes (Hara et al., 1994; Langlands et al., 2000; Schaefer et al., 2001). In Id1-HA-expressing cells, the endogenous Id1, but not the recombinant Id1-HA, was downregulated during this density-dependent differentiation process (Figure 4c, lower panel). Compared to wild-type and vector-transfected HaCaT cells, downregulation of endogenous Id1 was less pronounced in Id1-HA-expressing cells, possibly as a consequence of Id1-mediated inhibition of early differentiation (see below).
Id1 overexpression does not affect proliferation of HaCaT cells in monoculture
We next determined whether Id1 overexpression affects cell proliferation under exponential culture conditions in serum-containing medium and/or in response to TGF-β1 or activin A. Therefore, we performed 5-bromo-2′-deoxyuridine (BrdU) incorporation studies with different HaCaT/Id1-HA (n=5) and HaCaT/neo (n=3) clones. However, no significant difference in the proliferation rate of these cells was observed in monolayer culture (Figure 5a). Furthermore, we evaluated the effects of activin A and TGF-β1 on Id1-HA-expressing HaCaT and control cells. Figure 5b shows that the growth-inhibitory effect of TGF-β1 for keratinocytes (Shipley et al., 1986) was not abrogated by enhanced expression of Id1. Activin only showed a very weak antimitotic effect that was not influenced by Id1 overexpression.
Id1-overexpressing HaCaT cells differentiate abnormally in organotypic cultures
Finally, we determined if Id1 overexpression affects keratinocyte differentiation. Since HaCaT cells differentiate similarly to primary keratinocytes in three-dimensional organotypic cultures (Schoop et al., 1999; Maas-Szabowski et al., 2003), we established such cultures using either vector-transfected or HaCaT-Id1-HA cells. Figure 6 shows staining of cultures from one representative HaCaT/neo and one HaCaT/Id1-HA clone. In total, we tested five Id1-HA clones, and three clones of vector-transfected cells and we analysed two clones of each genotype in two independent experiments.
After 14 days of cultivation, vector-transfected HaCaT cells formed a multilayered, partially differentiated epidermis-like epithelium. After 4 weeks, the epithelium was fully differentiated and consisted of well-organized basal, spinous, and granular layers, and a parakeratotic cornified layer (Figure 6a). By contrast, the organotypic cocultures with Id1-HA-overexpressing HaCaT cells showed severe abnormalities. As shown in the hematoxylin/eosin-stained sections (Figure 6a), Id1-HA clones formed a hyperplastic epithelium after 2–4 weeks of culture. The time point of maximal hyperplasia varied among the Id1-HA cultures and was particularly early in cultures of the clone with the highest expression level (data not shown). Id1-HA cultures showed an increase in the number of viable cell layers, but a thinner cornified layer. Furthermore and in contrast to the control cultures, the Id1-HA cells had invaded the collagen gel, leading to a disorganized border between collagen gel and epithelium.
To verify the expression of Id1-HA in the epithelium of organotypic cultures, we performed immunofluorescence staining using an antibody directed against the HA-epitope. We found specific signals (pink nuclei) in cells of the basal and the suprabasal layers of cultures derived from HaCaT/Id1-HA clones (Figure 6b, left panels). Consistent with this expression pattern of Id1-HA, we found Id1 immunoreactivity in the basal layer of vector-transfected and Id1-HA-expressing cells, whereas suprabasal Id1-positive nuclei were only seen in Id1-HA cultures (Figure 6b, right panels).
In control cultures, the α6-integrin subunit is restricted to the basolateral side of basal keratinocytes (Figure 6c). In these cells it forms a dimer with the β4 subunit, and these α6β4 integrins are a component of hemidesmosomes that connect basal cells to the basement membrane (Watt, 2002). In Id1-HA-expressing cells, a more diffuse α6-integrin staining was observed, with staining around the basal cells and punctuate staining in a few suprabasal cells. A less-organized staining compared to control cells was also seen for the basement membrane marker laminin-5 (Figure 6d), further reflecting the disorganized basal cell layer and the invasion of cells into the collagen gel. No epithelial–mesenchymal transition occurred in the epithelium of Id1-HA cells as revealed by the lack of the mesenchymal marker protein vimentin in these cells (data not shown).
To analyse the degree of differentiation of Id1-overexpressing HaCaT cells in more detail, we stained sections with antibodies against characteristic early and late differentiation markers (shown for the 4-week cultures in Figure 6e and f). Whereas expression of the early differentiation marker keratin 10 (K10) started in the first suprabasal layer of control cultures, we found an irregular distribution of K10 in Id1-HA cultures (Figure 6e, and data not shown). In particular, the cells, which had invaded into the collagen gel, did not express K10. In addition, the number of K10-positive layers was also increased in most of the cultures. Most surprisingly, the late differentiation markers loricrin and filaggrin (Figure 6f and data not shown), which are characteristically expressed in the uppermost living cell layers, were present throughout the epithelium in a patchy distribution.
We next determined if the hyperplasia of the epidermal layer results in part from enhanced proliferation. As shown in Figure 6g, the proliferation rate of basal keratinocytes was slightly increased in Id1-HA cultures after 2 weeks (upper panel), but returned to normal levels after 4 weeks (lower panel). In all cultures, BrdU-positive cells were predominantly seen in the basal layer, although a few cells in the suprabasal layers of Id1-HA cultures had also incorporated BrdU.
Finally, we determined the rate of apoptosis in the different organotypic cultures by staining for cleaved caspase-3 (Figure 6h). As expected, we found caspase-3-positive cells in the most upper layer of all cultures, reflecting the disassembly of the cellular components during terminal differentiation. However, the number of apoptotic cells within the epithelium was strongly enhanced in 2-week and 4-week Id1-HA cultures. In particular, a large number of cells in the lower suprabasal layers stained positive for caspase-3 in these cultures.
Taken together, our results reveal that a small enhancement in the levels of Id1 affects proliferation, differentiation, and apoptosis of keratinocytes grown in organotypic cultures. This suggests that normal epidermis formation depends on a tight regulation of the levels of Id1.
Id proteins: novel targets of activin action in keratinocytes and fibroblasts
Activin is an important regulator of skin homeostasis and wound repair. However, there is little knowledge about the mechanisms underlying activin action in the skin, and the activin target genes are largely unknown. Here we show that activin suppresses the expression of id genes in keratinocytes in vitro. Most interestingly, downregulation of these genes was also seen in keratinocytes of activin-overexpressing transgenic mice, suggesting that id genes are also targets of activin action in vivo. This hypothesis is further supported by the inverse relationship between activin and Id expression in the hyperproliferative wound epidermis (Hübner et al., 1996, and this study), and indicates that endogenous activin also regulates the levels of Id proteins. Thus, in addition to TGF-β (Ling et al., 2002; Kretschmer et al., 2003; Siegel et al., 2003; Kowanetz et al., 2004), activin was identified as a novel regulator of Id gene expression.
Similar to TGF-β, activin has opposite effects on keratinocytes and fibroblasts. It slightly reduces keratinocyte proliferation (Shimizu et al., 1998; Seishima et al., 1999), but enhances growth of cultured fibroblasts (Ohga et al., 1996; Yamashita et al., 2004). The different responses of fibroblasts and keratinocytes to activin and TGF-β are reflected by the opposite regulation of Id expression in both cell types, suggesting that the up- or downregulation of Id proteins by activin/TGF-β could mediate – at least in part – the effects of these factors on proliferation of fibroblasts and keratinocytes. Consistent with this hypothesis, Id1 proteins are required for G1 progression of human fibroblasts (Hara et al., 1994), whereas overexpression of Id2 and Id3 in HaCaT cells abolished the antimitotic effect of TGF-β (Kowanetz et al., 2004).
Strong Id1 overexpression is detrimental for keratinocytes
To determine if Id1 affects keratinocyte proliferation, differentiation, and/or apoptosis, we generated stable HaCaT cell lines overexpressing Id1. Surprisingly, we only obtained cell lines that express the recombinant Id1-HA protein at similar or lower levels compared to the endogenous Id1. This finding is consistent with reports from other investigators who failed to overexpress Id1 in HaCaT cells (Langlands et al., 2000) and suggests that enhanced levels of this protein are detrimental. This hypothesis is further supported by our results, which revealed a higher rate of apoptosis of the Id1-overexpressing clones in organotypic cultures as well as by preliminary findings with the clone with the highest Id1 expression level. The latter showed a higher apoptotic rate in response to staurosporine treatment or UVB irradiation, and the high-expressing cells in this culture were lost upon passaging (data not shown). Consistent with our findings, stable transfection of rat embryo fibroblasts with Id2 resulted in apoptosis (Norton et al., 1998). Therefore, the extent of Id overexpression that can be achieved seems to be limited.
Id1 is not required for the growth-suppressing effect of TGF-β1
Although the extent of Id1-HA overexpression was minor under normal culture conditions, significant overexpression was observed under conditions where the endogenous protein was downregulated, such as in response to TGF-β1 or upon induction of differentiation. Therefore, these cell lines were suitable to determine the role of Id1 downregulation in the growth suppressive effect of TGF-β and activin. However, forced expression of Id1-HA in HaCaT cells did not significantly affect the proliferation rate of keratinocytes in serum-containing medium and in response to activin and TGF-β1. This finding is consistent with results from Kang et al. (2003), who demonstrated that transient overexpression of Id1 in HaCaT cells does not diminish the TGF-β cytostatic response. However, different results were obtained with overexpressed Id2 and Id3 in HaCaT cells. In this case, the growth-inhibiting effect of TGF-β was abrogated (Kowanetz et al., 2004). In addition to the different regulation in cultured cells and skin wounds, this finding suggests different functions of Id1 as compared to the other Id proteins.
A novel role of Id1 in epidermal homeostasis
In addition to their effects on proliferation, Id proteins potently suppress differentiation of various cell types (Ruzinova and Benezra, 2003). Therefore, we tested if Id1 also affects differentiation of keratinocytes. Since full differentiation of HaCaT cells can only be achieved in three-dimensional organotypic cultures (Schoop et al., 1999), we used this technology to determine the effect of Id1 overexpression on keratinocyte differentiation. Indeed, the epithelia formed by Id1-HA-expressing cells were hyperthickened and disorganized, and under these conditions, Id1 overexpression transiently enhanced the proliferation rate of keratinocytes. Thus, fibroblast-derived paracrine acting factors and/or the three-dimensional architecture are obviously required for Id1-mediated stimulation of keratinocyte proliferation. Interestingly, enhanced levels of Id1 were also seen in the hyperplastic epidermis of psoriatic patients (Bjorntorp et al., 2003), suggesting a functional role of Id1 in the pathogenesis of this disease. In addition, high levels of Id proteins in the hyperproliferative wound epidermis may enhance proliferation and prevent premature onset of early differentiation of wound keratinocytes, thereby contributing to wound re-epithelialization. In suprabasal cells of the wound epidermis, high levels of activin and/or TGF-β (Frank et al., 1996; Hübner et al., 1996) are likely to suppress Id expression, resulting in growth inhibition and induction of differentiation of these cells.
During wound healing, keratinocytes from the border of the lesion detach from the basement membrane and repair the defect by transforming from a sessile into a mobile phenotype (Martin, 1997). Schaefer et al. (2001) observed upregulation of Id1 in keratinocytes upon detachment from their growth substratum. This and several reports on the crucial role of Id overexpression in tumor invasiveness and metastasis (Lin et al., 1999) suggest an important influence of Id proteins not only on proliferation, but also on the stimulation of cell motility after skin injury.
We also found strong abnormalities in the differentiation pattern as well as in the rate of apoptosis of Id1-HA cell cultures. Thus, our results demonstrate that enhanced levels of Id1 disturb the balance between keratinocyte proliferation and differentiation/apoptosis, resulting in a hyperplastic, disorganized epidermis. These findings suggest that a tight regulation of the levels of this transcriptional regulator is required for epidermal homeostasis. Finally, our findings also suggest that Id1 induces invasive growth of keratinocytes, since we observed a rather extensive ingrowth of Id1-HA-expressing HaCaT cells into the dermal equivalent. As this phenotype is not unique for Id1 but is also observed with other HaCaT cell clones, which express genes that alter the differentiation process (PB and SW, unpublished data), it is tempting to speculate that with the altered differentiation, a program is induced that causes activation of collagen-degrading enzymes and thus allows invasion into the dermal equivalent. Future experiments will address whether regulation of the expression of specific proteases such as collagenases (matrix metalloproteases) or cathepsins may be another as yet unknown activity of Id1 in keratinocytes.
Materials and methods
Cell culture and growth factor treatment of keratinocytes
HaCaT keratinocytes (Boukamp et al., 1988) were grown to confluence in Dulbecco's modified Eagle's medium (DMEM) (Sigma, Deisenhofen, Germany) containing 1% penicillin/streptomycin and 10% fetal calf serum (FCS) (BioConcept Amimed, Allschwil, Switzerland). They were rendered quiescent by serum starvation for 24–32 h and subsequently treated with fresh DMEM containing 1 ng/ml TGF-β1 (R&D Systems, Wiesbaden, Germany) or 18 ng/ml activin A (R&D Systems). These activin concentrations were reported to reduce keratinocyte proliferation and to induce their differentiation (Seishima et al., 1999), and they elicit maximal transcriptional responses in HaCaT cells (Werner et al., 2001). Cells were harvested at different time points after addition of the growth factor and used for RNA isolation or preparation of cell lysates.
Primary human dermal fibroblasts from foreskin biopsies were cultured in DMEM containing 1% penicillin/streptomycin and 10% FCS and treated with growth factors as described for HaCaT cells. Human dermal fibroblasts from adult trunk skin were isolated from explant cultures of de-epidermized dermis. They were expanded up to passage 3–4 in DMEM/10% FCS and cryo-preserved until used in organotypic cultures.
The preparation of the dermal equivalent based on type I collagen hydrogels has been described previously (Smola et al., 1998; Stark et al., 1999; Maas-Szabowski et al., 2003). Briefly, 80% (vol.) type I collagen in 0.1% acetic acid (4 mg/ml; rat tail tendon) were mixed on ice with 10% (vol.) of 10 × Hank's buffered saline containing phenol red. After neutralizing with 2 M NaOH, a fibroblast suspension in FCS was added under stirring (1 × 105 cells/ml, final concentration). This mixture was transferred into membrane filter inserts (Falcon no. 3901, 3 μm pores, BD-Biosciences, Heidelberg, Germany), placed in deep well plates (BD-Biosciences no. 5467) and gelled at 37°C. Glass rings were placed on the gels to allow keratinocyte seeding. The gels were equilibrated for 24 h in FAD medium with 10% FCS, 10−10 M cholera toxin, 0.4 μg hydrocortisone, and 50 μg L-ascorbic acid (Sigma) per ml. Cocultures were obtained by seeding 1 × 106 keratinocytes inside the glass rings (3 × 105 cells per cm2). After submersed incubation overnight, the glass rings were removed and the medium level was lowered to the base of the dermal equivalent, thereby exposing the culture surface to the air-medium interface. Cultivation was continued for 2 or 4 weeks in FAD medium with 10% FCS, 10−10 M cholera toxin, 0.4 μg hydrocortisone, and 50 μg L-ascorbic acid/ml with medium change every other day.
Hybridization of cDNA arrays
Polyadenylated RNA was prepared from total cellular RNA using the mRNA Separator Kit (Clontech Laboratories, Palo Alto, CA, USA) following the instructions of the manufacturer. The poly(A)+-mRNA was isolated by affinity chromatography using oligo-d(T)-cellulose columns. The Atlas human cDNA expression array I (Clontech) was hybridized with radioactively labeled cDNA from quiescent and activin A-treated HaCaT cells according to the manufacturer's instructions.
Generation of HaCaT cells stably expressing Id1-HA
The full-length human id1 cDNA was amplified from HaCaT cDNA using RT–PCR. In an additional PCR step, an in-frame HA epitope was inserted in front of the stop codon of the id1 cDNA. The construct was subcloned into the expression vector pIRES/neo (Clontech). For stable transfection, HaCaT cells were seeded at 50–70% confluence and transfected using LipofectAMINE (Invitrogen, Basel, Switzerland) according to the manufacturer's instructions. Cells stably transfected with the pIRES/neo expression vector were selected by cultivation in growth medium containing 400 mg/l G418 (Invitrogen).
RNA isolation and RPA
Isolation of total cellular RNA was performed as described by Chomczynski and Sacchi (1987). RPAs were carried out according to Werner et al. (1993). The following cDNA templates were amplified by RT–PCR using RNA from mouse wounds or HaCaT keratinocytes: a human id1 cDNA corresponding to nucleotides (nt) 121–365 of the published sequence (Hara et al., 1994), a human id2 cDNA (nt 96–308 of the cDNA; Hara et al., 1994), a human id3 cDNA (nt 738–936 of the cDNA; Deed et al., 1994), a mouse id1 cDNA (nt 251–528 of the cDNA; NM_010495), a mouse id2 cDNA (nt 1119–1349 of the cDNA; AF077860), and a mouse id3 cDNA (nt 57–255 of the cDNA; NM_010813). These cDNAs were cloned into the transcription vector pBluescript KSII(+) (Stratagene, La Jolla, CA, USA). As a loading control, either 1 μg of the RNA samples was loaded on a 1% agarose gel prior to hybridization and stained with ethidium bromide or the RNA was hybridized with a probe for murine glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (nt 566–685 of the cDNA; NM_008048.1) or with a probe for human GAPDH (nt 580–695 of the cDNA; BC_001601). Autoradiograms were scanned and analysed with the Scion Image software (Scion Corporation, MD, USA).
Preparation of protein lysates from HaCaT cells and fibroblasts
HaCaT cells were lysed in urea buffer (10 mM Tris [pH 8.0], 9.5 M urea, 2 mM EDTA, 2 mM PMSF, 1 mM dithiothreitol). Lysates were sonicated and centrifuged for 15 min at 4°C. Protein concentrations in the supernatant were determined using the BCA Protein Assay Reagent (Pierce, Rockford, IL, USA).
Western blot analysis
Proteins were separated by SDS polyacrylamide gel electrophoresis and transferred to a nitrocellulose membrane. Antibody incubations were performed in 3% non-fat dry milk in TBS-T (10 mM Tris-HCl pH7.4, 150 mM NaCl, 0.1% Tween 20). The following antibodies were used: a rabbit polyclonal antibody against Id1 (Santa Cruz Biotechnology, Santa Cruz, CA, USA), a rabbit polyclonal antibody against the HA epitope (Santa Cruz), and a mouse monoclonal antibody against β-actin (Sigma). Detection was performed with the enhanced chemoluminescence detection system (ECL; Amersham).
Immunofluorescence and histological analysis
Tail skin from BALB/c and CD1 mice and wounds from BALB/c mice were frozen in tissue freezing medium (Jung, Nussloch, Germany). Acetone-fixed frozen sections (6 μm) were incubated overnight at 4°C with rabbit polyclonal antibodies directed against Id1, Id2, or Id3 (Santa Cruz) diluted in PBS containing 12% BSA, 0.1% NP-40, and 0.1% Tween 20. After three washes with PBS/0.1% Tween 20, sections were incubated for 1 h with an anti-rabbit-Cy3 secondary antibody (Jackson ImmunoResearch Laboratories, West Grove, PA, USA), washed again and mounted with Mowiol (Hoechst, Frankfurt, Germany). Organotypic cultures were fixed at 4°C in acidic ethanol and embedded in paraffin. Alternatively, they were frozen in tissue-freezing medium without prior fixation. Paraffin sections (7 μm) were de-waxed and incubated overnight at 4°C with a mouse monoclonal antibody directed against keratin 10 (Dako), or rabbit polyclonal antibodies directed against keratin 14, filaggrin, loricrin (all from BabCO, Richmond, CA, USA), and the HA-epitope (Santa Cruz). Frozen sections were fixed with ice-cold acetone or 4% paraformaldehyde and incubated with the Id1 antibody described above, a rat monoclonal antibody directed against α6-integrin (BD Bioscience PharMingen, San Diego, CA, USA), or rabbit polyclonal antibodies directed against the γ2-chain of laminin-5 (kindly provided by Dr Takako Sasaki, Martinsried, Germany) or cleaved caspase-3 (Cell Signaling Technology, Beverly, MA, USA). As secondary antibodies, anti-rabbit-Cy3 (red), anti-rat-Cy3, and anti-mouse-Cy2 (green) (Jackson ImmunoResearch) were used. Sections were counterstained with DAPI (1 μg/ml) and photographed using a Zeiss Axioplan fluorescence microscope. For histological analysis, paraffin-embedded OTCs were sectioned, dewaxed and stained with hematoxylin/eosin according to standard procedures.
BrdU incorporation studies
1.2 × 105 HaCaT cells were seeded into 3.5 cm dishes. After 24 h of cultivation, cells were treated with 18 or 90 ng/ml activin A or 1 ng/ml TGF-β1 for 24 h and incubated with BrdU (Sigma, 100 μ M) for 2 h before fixation with 4% PFA. To permeabilize the cell membrane and to denature the DNA, cells were treated with 2 M HCl/0.1% Triton X-100 for 30 min. Subsequently, 0.1 M Na-borate (pH 8.5) was added for 5 min. Immunofluorescence staining of BrdU-positive cells was performed by overnight incubation at 4°C with an FITC-conjugated monoclonal antibody directed against BrdU (Roche, Rotkreuz, Switzerland). Cells were counterstained with propidium iodide (2 μg/ml), mounted, and photographed using a Zeiss AxioCam HRc camera. For proliferation analysis, the ratio of total cell number to the number of BrdU-positive cells was determined by counting cells using Openlab software (GraphPad Software Inc, San Diego, USA).
Transgenic activin βA-overexpressing CD1 mice were described previously (Munz et al., 1999). BALB/c mice were obtained from RCC (Füllinsdorf, Switzerland). Mice were housed and fed according to federal guidelines.
Wounding and preparation of wound tissue
Mice were anesthetized by intraperitoneal injection of ketamine/xylazine. Two full-thickness excisional wounds, 5 mm in diameter, were made on either side of the dorsal midline as described previously (Werner et al., 1994). Wounds were left uncovered and harvested at different time points after injury. For RPA, the complete wounds including 2 mm of the epithelial margins were excised and immediately frozen in liquid nitrogen until used for RNA isolation. Nonwounded back skin served as a control. For immunofluorescence analyses, the complete wounds were isolated, bisected, and frozen in tissue-freezing medium. All experiments with mice were performed with permission from the local veterinary authorities of Zurich, Switzerland.
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We thank Christiane Born-Berclaz and Iris Nord for excellent technical assistance and Dr Takako Sasaki, Max-Planck-Institute of Biochemistry, Martinsried, Germany, for kindly providing the laminin-5 antibody. This work was supported by a grant from the Swiss National Science Foundation (31-61358.00) to SW and a grant from the EC/Swiss Ministry for Education and Research (LSHG-CT-2003-503447, WOUND) to SW and PB.
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