In two-stage skin chemical carcinogenesis, phorbol ester 12-O-tetradecanoylphorbol-13-acetate (TPA) acts as a promoter essential for clonal expansion of the initiated cells carrying the activated ras oncogenes. Although protein kinase C (PKC) isozymes are the main targets of TPA, their role in tumor promotion remains controversial. We previously reported that mice lacking a Ras/Rap effector phospholipase Cε (PLCε−/− mice) exhibited marked resistance to tumor formation in the two-stage skin carcinogenesis. PLCε−/− mice also failed to exhibit basal layer cell proliferation and epidermal hyperplasia induced by TPA, suggesting a role of PLCε in tumor promotion. Here, we show that PLCε−/− mice exhibit resistance to TPA-induced skin inflammation as assessed by reduction in edema, granulocyte infiltration, and expression of a proinflammatory cytokine, interleukin-1α (IL-1α). On the other hand, the proliferative potentials of keratinocytes or dermal fibroblasts in culture remain unaffected by the PLCε background, suggesting that the PLCε's role in tumor promotion may be ascribed to augmentation of inflammatory responses. In dermal fibroblast primary culture, TPA can induce activation of the PLCε lipase activity, which leads to the induction of IL-1α expression. Experiments using small interfering RNA–mediated knockdown indicate that this activation is mediated by Rap1, which is activated by a TPA-responsive guanine nucleotide exchange factor RasGRP3. Moreover, TPA-induced activation of Rap1 and PLCε is inhibited by a PKC inhibitor GF109203X, indicating a crucial role of PKC in signaling from TPA to PLCε. These results imply that two TPA targets, RasGRP3 and PKC, are involved in TPA-induced inflammation through PLCε activation, leading to tumor promotion. [Cancer Res 2008;68(1):64–72]

Phosphoinositide-specific phospholipase C (PLC) plays pivotal roles in intracellular signaling by catalyzing the hydrolysis of phosphatidylinositol 4,5-bisphosphate into two important second messengers, diacylglycerol (DAG), and inositol 1,4,5-trisphosphate (IP3). DAG binds to a number of its target proteins, represented by protein kinase C (PKC) isozymes, and regulates their activities. IP3 opens calcium channels on the surface of the intracellular stores, causing increase in the cytosolic free calcium concentration. At least 13 mammalian PLC isoforms have been identified and organized into six classes (β, γ, δ, ε, ζ, and η) based upon the similarities in their structures and regulatory mechanisms (1, 2).

We and others identified PLCε as an effector of Ras family small GTPases (Ras, Rap1, and Rap2), which bind directly to its Ras-associating domain (36). Subsequent studies have shown that PLCε is also activated by small GTPase RhoA and heterotrimeric G proteins Gα12 and Gβ1γ2 (6). These multiple regulatory mechanisms enable PLCε to mediate signals from a wide variety of cell surface receptors, including receptor tyrosine kinases (7) and G protein–coupled receptors (8). In addition, PLCε functions as a guanine nucleotide exchange factor (GEF) for Rap1 by its CDC25 homology domain (9). The physiologic role of PLCε has been intensively studied using various animals carrying artificial or spontaneous mutations in its chromosomal gene. Positional cloning of the gene responsible for a nephrotic syndrome identified mutations in the human PLCε gene PLCE1 (10). Knockdown of the PLCε orthologue in the zebrafish resulted in loss of the filtration barrier in the glomerular podocytes resembling the human symptom (10). Mice homozygous for the functionally inactivated PLCε allele (PLCε−/− mice) exhibited semilunar valvulogenesis defect, leading to cardiac dilation (11), and mice with total disruption of the PLCε gene developed cardiac hypertrophy under an extreme cardiac stress (12). Targeted inactivation of the PLCε orthologue in the nematode Caenorhabditis elegans resulted in delayed dilation of the spermatheca-uterine valve, leading to defective ovulation (13).

We showed that PLCε−/− mice exhibited marked resistance to tumor formation in the two-stage skin chemical carcinogenesis protocol using 7,12-dimethylbenz(a)anthracene (DMBA) as an initiator and a phorbol ester, 12-O-tetradecanoylphorbol-13-acetate (TPA), as a promoter (14). PLCε−/− mice also failed to exhibit basal layer cell proliferation and epidermal hyperplasia induced by TPA, suggesting a role of PLCε in tumor promotion. As TPA is a molecular mimic of DAG, a variety of DAG target proteins carrying the C1 domains are potential effectors for the TPA-dependent tumor promotion, such as conventional PKCs, novel PKCs, protein kinase D, α and β chimaerins, Munc13-1∼4, diacylglycerol kinases β and γ, and RasGRP1∼4 (15). RasGRPs, also called CalDAG-GEFs, are specific GEFs for Ras family small GTPases (16). The association with DAG/TPA is sufficient for activation of chimaerins, whereas phosphorylation by PKC is also required for activation of RasGRP3 (15). Although PKC isozymes have been most intensively investigated, their role in de novo carcinogenesis, including tumor promotion, remains controversial. For instance, overexpression of PKCβII in the colon resulted in hyperproliferation and increased sensitivity to carcinogen-induced cancer (17). In contrast, transgenic overexpression of α or δ isozyme of PKC exhibited inhibitory effects on tumor promotion with TPA (18, 19), and targeted disruption of the PKCα or PKCη gene resulted in increased tumor formation (2022), indicating that these isozymes may function as tumor suppressors. Such observations led to the reassessment of the role of PKCs and suggested the involvement of non-PKC molecules in tumor promotion (15).

The causal relationship between tumor promotion and inflammation, an old hypothesis, has gained substantial experimental supports from recent studies (23). However, molecular and cellular mechanisms mediating this relationship remain unsolved. In this study, we show that PLCε plays a crucial role in TPA-induced inflammation, which seems to account for its role in TPA-dependent tumor promotion. PLCε is activated by TPA in dermal fibroblasts, which is mediated by RasGRP3-dependent activation of Rap1. Interestingly, PKC activation, too, is required for this Rap1 activation, the mechanism of which will be discussed.

Animals. Mice carrying the inactivated PLCε allele (PLCε), created by in-frame deletion of an exon coding for the catalytic X domain, were generated as described previously (11, 14). They were backcrossed to C57BL/6 strain eight times. PLCε+/+ and PLCε−/− littermates were produced by cross-breeding of PLCε+/− mice and were used for the experiments. All the animals were maintained at the animal facilities of Kobe University Graduate School of Medicine. ICR mice were purchased from Japan SLC, Inc. The use and care of the animals were reviewed and approved by the Institutional Animal Committee of Kobe University Graduate School of Medicine.

Antibodies and chemicals. Primary antibodies used here were antimouse keratin 1 (K1; Covance), antimouse keratin 14 (K14; Covance), anti-Rap1 (sc-65; Santa Cruz Biotechnology, Inc.), anti-actin (sc-8432; Santa Cruz Biotechnology, Inc.), anti-mouse Gr-1 (MAB1037; R&D Systems, Inc.), anti–mitogen-activated protein kinase (MAPK; 9102; Cell Signaling Technology, Inc.), anti–phosphorylated MAPK (9106; Cell Signaling Technology, Inc.), anti–Pan-Ras (OP21; Calbiochem), and antivimentin (ab7783; Abcom). Anti-PLCε antibody raised against the C terminus of mouse PLCε was described (24). Secondary antibodies conjugated with horseradish peroxidase were purchased from GE Healthcare. Fluorescently active secondary antibodies labeled with Alexa Fluor 488 or Alexa Fluor 546 were purchased from Invitrogen. A PLC inhibitor, U73122, and a PKC inhibitor, GF109203X, were purchased from Calbiochem.

Preparation and culture of skin keratinocytes and fibroblasts. Epidermal keratinocytes and dermal fibroblasts were isolated from the dorsal skin of 1-day-old to 3-day-old newborn mice and cultured as described previously (25). The purity of the cells was over 99% as assessed by immunostaining with anti-K14 and anti-vimentin antibodies for keratinocytes and fibroblasts, respectively. For organotypic culture of keratinocytes, dermal fibroblasts isolated from wild-type mice were used for reconstitution of the dermal equivalent as described (25).

Preparation of splenic B lymphocytes and non–B lymphocytes and peritoneal macrophages. Splenic B lymphocytes and non–B lymphocytes were prepared as described (26). Thioglycollate-elicited peritoneal macrophages were prepared according to the standard procedure. Briefly, mice were i.p. injected with 3 mL of sterile 4% thioglycollate (Sigma). Four days later, the peritoneal lavage was collected and, after removal of fibroblasts through adherence onto glass plates, used as a macrophage preparation.

Assessment of TPA-induced skin responses. The backside of the ears of 8-week-old to 12-week-old male mice was topically treated with 50 μL of 50 μg/mL TPA (P-8139; Sigma) dissolved in acetone. Ear thickness was measured with calipers, and ear swelling was calculated as (thickness at each time point) − (thickness at 0 h). Histologic analysis with H&E staining and immunohistochemical analysis of the skin sections were carried out essentially as described (11, 14).

Reverse transcription–PCR analysis. Reverse transcription–PCR (RT-PCR) was performed as described previously (11, 24). The sequences of the primers used for PCR are listed in Supplementary Table S1.

Western blot analysis. Cells were solubilized in lysis buffer [50 mmol/L Tris-Cl (pH 7.5), 250 mmol/L NaCl, 1 mmol/L EDTA, 0.5% (v/v) Triton X-100, 1 mmol/L phenylmethylsulfonyl fluoride (PMSF), 10 mmol/L NaF, 1 mmol/L Na3VO4, 20 mmol/L β-glycerophosphate, 1 μmol/L leupeptin]. SDS-PAGE and immunoblotting were performed as described previously (24).

Measurements of Ras-GTP and Rap1-GTP. The cellular levels of Ras-GTP and Rap1-GTP were determined by pull-down assays using glutathione S-transferase (GST) fusions of Raf-1 Ras-binding domain (GST–Raf-1–RBD) and RalGDS Ras-interacting domain (GST-RalGDS-RID), respectively (27). After serum-starving in 0.1% FCS overnight, cells were stimulated with TPA or vehicle for 5 min and lysed in magnesium lysis buffer [50 mmol/L Tris-HCl (pH 7.4), 150 mmol/L NaCl, 0.5% Nonidet P-40, 20 mmol/L MgCl2, 1 μmol/L leupeptin, 1 mmol/L PMSF], and soluble fractions were collected by centrifugation. Protein concentrations were determined by Bradford method, and 0.7 and 1 mg of the lysate protein were used for pull-down of Ras-GTP and Rap1-GTP, respectively, which were detected by Western blotting with anti–Pan-Ras and anti-Rap1 antibodies, respectively.

Measurement of cytosolic free calcium concentration. Cytosolic free calcium concentration was monitored in cells loaded with acetoxymethyl ester form of Fura-2 (Fura-2 AM; ref. 28). Dermal fibroblasts in culture were collected by trypsinization and washed once with DMEM containing 10% FCS and once with solution A [5.4 mmol/L KCl, 0.3 mmol/L Na2HPO4, 0.4 mmol/L KH2PO4, 4.2 mmol/L NaHCO3, 1.3 mmol/L CaCl2, 0.5 mmol/L MgCl2, 0.6 mmol/L MgSO4, 1.4 mmol/L NaCl, 5.6 mmol/L d-glucose, 10 mmol/L HEPES (pH 7.4), 200 μmol/L sulfinpyrazone (S-9509; Sigma)]. Cells were incubated for 30 min at 37°C in solution A containing 5 μmol/L Fura-2 AM (Nacalai Tesque, Inc.) and 0.02% (v/v) Plaronic F27 (P-3000MP; Molecular Probes). After washing twice with solution A, the cells were suspended at 7.5 × 105 cells/mL in solution A containing 0.1% (w/v) bovine serum albumin and subjected to measurement of the emission at 510 nm with the excitation at 340/380 nm using F-4500 fluorometer (Hitachi). Fura-2 loading was confirmed by subsequent treatment of the cells with 10 μmol/L ionomycin followed with 0.1% (v/v) Triton X-100. Results are expressed as relative concentrations of cytosolic free calcium calculated as described (29).

Gene silencing by small interfering RNA. Dermal fibroblasts (4 × 106 cells) suspended in 1 mL of OptiMEM (Invitrogen) were transfected with 600 nmol/L Stealth small interfering RNAs (siRNA; Invitrogen) using GenePulser (Bio-Rad) at 400 μF, 380 V. After electroporation, cells were cultured in DMEM containing 10% FCS for 48 h and subjected to further experiments. The identification numbers of the Stealth siRNAs used were MSS216065 and MSS216067 for RasGRP3 (abbreviated as grp65 and grp67, respectively), MSS201244, and MSS201242 for Rap1A (a44 and a42, respectively), and MSS210941 and MSS210942 for Rap1B (b41 and b42, respectively). Stealth RNA interference negative control kit with Low GC or Med GC (Invitrogen) was used as negative control.

Statistical analysis. Values are expressed as the averages ± SDs. The unpaired Student's t test was performed for determination of P values.

Proliferative potentials of skin keratinocytes and fibroblasts are not affected by the PLCε background. Our previous observation on the crucial roles of PLCε in two-stage skin carcinogenesis and TPA-induced epidermal hyperplasia prompted us to examine its role in proliferation of skin cells in culture. We first analyzed the expression levels of PLCε in primary cultured epidermal keratinocytes and dermal fibroblasts. The keratinocytes in culture were positive for K14 and represented the proliferative populations residing in the basal cell layer of the epidermis. RT-PCR analysis showed that PLCε is expressed much more abundantly in fibroblasts than in keratinocytes (Fig. 1A). Western blot analysis using anti-PLCε antibody detected two immunoreactive bands, which presumably corresponded to the splicing variants PLCε1a and PLCε1b (12), and showed more abundant expression in fibroblasts (Fig. 1A). These data are consistent with our previous immunohistochemical observation that PLCε is expressed more weakly in K14-positive proliferative keratinocytes than in K1-positive differentiating keratinocytes (ref. 14; also see Supplementary Fig. S1A). We next evaluated the proliferative potential of keratinocytes and fibroblasts established from PLCε+/+ and PLCε−/− mice. We used the organotypic culture system, in which keratinocytes were seeded onto the dermal equivalent consisting of collagen fibers and dermal fibroblasts isolated from wild-type mice and induced to differentiate by air exposure and raise of calcium concentration in the culture medium (25). Essentially, no difference was observed in their proliferation, as indicated by the number of layers negative for K1, as well as in their differentiation, as estimated from the number of cells positive for K1 (Fig. 1B). Likewise, growth rate of dermal fibroblasts cultured in the presence of 10% FCS was not affected by the PLCε background (Fig. 1C). Furthermore, neither MAPK activation by TPA and epidermal growth factor (EGF; Fig. 1D,, top) nor c-fos induction by TPA (Fig. 1D , bottom) was altered in dermal fibroblasts depending on the PLCε background, suggesting that PLCε did not affect these growth-promoting signaling pathways. Thus, PLCε did not seem to directly affect the proliferation of epidermal keratinocytes or dermal fibroblasts per se.

Figure 1.

Effect of PLCε knockout in proliferation and differentiation of skin cells in culture. A, expression of PLCε. The expression level of PLCε in keratinocytes and fibroblasts cultured from the dorsal skin of newborn ICR mice was analyzed by RT-PCR (left) and Western blotting (right). For RT-PCR, reverse transcriptase (RTase) minus controls and internal controls [glyceraldehyde-3-phosphate dehydrogenase (GAPDH)] are shown. Cell lysates (30 μg of protein) were fractionated by SDS-PAGE and subjected to Western blotting with anti-PLCε antibody. Antiactin blot was used as loading controls. Arrow heads, bands corresponding to PLCε1a and PLCε1b. B, proliferation and differentiation of keratinocytes and fibroblasts in culture. Epidermal keratinocytes isolated from the dorsal skins of PLCε+/+ (+/+) and PLCε−/− (−/−) newborn mice were subjected to the organotypic culture. After induction of keratinocyte differentiation for 4 and 7 d, cross-sections of the culture were stained with anti-K1 antibody (green) and 4′,6-diamidino-2-phenylindole (DAPI, blue, left). Dermal fibroblasts were seeded at the density of 1 × 105 cells per Φ 60-mm plate and cultured in the presence of 10% FCS. The cell numbers were determined from three plates in triplicate at each time point using a hematocytometer; points, average; bars, SD (right). C, MAPK activation (top) and c-fos induction (bottom) in dermal fibroblasts from PLCε+/+ and PLCε−/− mice. Dermal fibroblasts were stimulated with TPA (100 ng/mL) or EGF (20 ng/mL) for 5 min and subjected to Western blotting with anti–phosphorylated MAPK antibody or anti-MAPK antibody. The level of c-fos mRNA was examined by RT-PCR after treatment of the cells with the indicated concentrations of TPA for 2 or 6 h.

Figure 1.

Effect of PLCε knockout in proliferation and differentiation of skin cells in culture. A, expression of PLCε. The expression level of PLCε in keratinocytes and fibroblasts cultured from the dorsal skin of newborn ICR mice was analyzed by RT-PCR (left) and Western blotting (right). For RT-PCR, reverse transcriptase (RTase) minus controls and internal controls [glyceraldehyde-3-phosphate dehydrogenase (GAPDH)] are shown. Cell lysates (30 μg of protein) were fractionated by SDS-PAGE and subjected to Western blotting with anti-PLCε antibody. Antiactin blot was used as loading controls. Arrow heads, bands corresponding to PLCε1a and PLCε1b. B, proliferation and differentiation of keratinocytes and fibroblasts in culture. Epidermal keratinocytes isolated from the dorsal skins of PLCε+/+ (+/+) and PLCε−/− (−/−) newborn mice were subjected to the organotypic culture. After induction of keratinocyte differentiation for 4 and 7 d, cross-sections of the culture were stained with anti-K1 antibody (green) and 4′,6-diamidino-2-phenylindole (DAPI, blue, left). Dermal fibroblasts were seeded at the density of 1 × 105 cells per Φ 60-mm plate and cultured in the presence of 10% FCS. The cell numbers were determined from three plates in triplicate at each time point using a hematocytometer; points, average; bars, SD (right). C, MAPK activation (top) and c-fos induction (bottom) in dermal fibroblasts from PLCε+/+ and PLCε−/− mice. Dermal fibroblasts were stimulated with TPA (100 ng/mL) or EGF (20 ng/mL) for 5 min and subjected to Western blotting with anti–phosphorylated MAPK antibody or anti-MAPK antibody. The level of c-fos mRNA was examined by RT-PCR after treatment of the cells with the indicated concentrations of TPA for 2 or 6 h.

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TPA-induced inflammation is attenuated in PLCε−/− skin. TPA treatment is known to induce inflammation in the skin, and a critical role of inflammation in tumor promotion has been widely acknowledged (23). During the chemical carcinogenesis experiments, we had noticed that TPA-induced edema in the dorsal skin, which seemed several hours after application, was considerably weaker in PLCε−/− mice compared with PLCε+/+ mice (data not shown). Here, we quantitatively examined the extent of the skin edema by treating the ears with TPA and measuring their thickness. Histologic analysis of the ears, prepared 10 h after TPA application, revealed marked spongiosis and extensive infiltration of leukocytes in the edematous dermis of PLCε+/+ mice. However, the extent of such changes was substantially reduced in PLCε−/− mice (Fig. 2A). Examination of the time course of the ear swelling in PLCε+/+ mice showed that the swelling reached a peak around 9 to 12 h after TPA application and started partial resolution thereafter (Fig. 2B). In contrast, the swelling in PLCε−/− mice showed a plateau with a moderate level around 9 to 24 h, suggesting that the early peak phase of the edema, observed in PLCε+/+, was markedly attenuated (Fig. 2B). Calculation of the number of infiltrated granulocytes in the ear sections 10 h after TPA application indicated that TPA-induced infiltration of inflammatory leukocytes was significantly reduced in PLCε−/− mice compared with PLCε+/+ mice (Fig. 2C). These results indicated that TPA-induced inflammation was attenuated in PLCε−/− mice, suggesting the crucial role of PLCε in augmentation of inflammatory responses.

Figure 2.

Comparison of TPA-induced inflammatory responses between PLCε+/+ and PLCε−/− mice. A, measurement of TPA-induced ear swelling. Ears of male mice were treated with TPA or acetone vehicle. At 10 h after application of TPA or acetone vehicle, the ears of PLCε+/+ and PLCε−/− mice were collected, and their cross-sections were subjected to H&E staining (left). The thickness of the ear of each mouse was determined by averaging the values measured at 9 or 10 independent regions of the cross-section. Black dots, thickness of each mouse; red lines, average of four mice of each genotype (right). Bar, 200 μm. **, P < 0.01 between PLCε+/+ and PLCε−/− mice. B, time course of TPA-induced ear swelling. The thickness of the TPA-treated ears of three PLCε+/+ and three PLCε−/− mice was measured at the indicated time points. The extent of ear swelling was calculated as described in Materials and Methods; points, average; bars, SD. **, P < 0.01 between PLCε+/+ and PLCε−/− mice at each time point. C, measurement of the densities of infiltrated granulocytes. The density of granulocytes of the ear of each mouse was determined by averaging the granulocyte counts in 15 to 30 randomly chosen regions of the ear section used in A under high magnification (left). Black dots, value of each mouse; red lines, average value of four mice of each genotype (right). Bar, 50 μm. *, P < 0.05 between PLCε+/+ and PLCε−/− mice.

Figure 2.

Comparison of TPA-induced inflammatory responses between PLCε+/+ and PLCε−/− mice. A, measurement of TPA-induced ear swelling. Ears of male mice were treated with TPA or acetone vehicle. At 10 h after application of TPA or acetone vehicle, the ears of PLCε+/+ and PLCε−/− mice were collected, and their cross-sections were subjected to H&E staining (left). The thickness of the ear of each mouse was determined by averaging the values measured at 9 or 10 independent regions of the cross-section. Black dots, thickness of each mouse; red lines, average of four mice of each genotype (right). Bar, 200 μm. **, P < 0.01 between PLCε+/+ and PLCε−/− mice. B, time course of TPA-induced ear swelling. The thickness of the TPA-treated ears of three PLCε+/+ and three PLCε−/− mice was measured at the indicated time points. The extent of ear swelling was calculated as described in Materials and Methods; points, average; bars, SD. **, P < 0.01 between PLCε+/+ and PLCε−/− mice at each time point. C, measurement of the densities of infiltrated granulocytes. The density of granulocytes of the ear of each mouse was determined by averaging the granulocyte counts in 15 to 30 randomly chosen regions of the ear section used in A under high magnification (left). Black dots, value of each mouse; red lines, average value of four mice of each genotype (right). Bar, 50 μm. *, P < 0.05 between PLCε+/+ and PLCε−/− mice.

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Role of PLCε in TPA-induced expression of proinflammatory molecules. Attenuated inflammation in PLCε−/− mice prompted us to examine the expression of proinflammatory molecules at the early stage of TPA treatment. We analyzed the expression levels of representative proinflammatory molecules on the ear at 3 and 6 h after TPA application by semiquantitative RT-PCR (Fig. 3A). We found that TPA-induced up-regulation of interleukin-1α (IL-1α) mRNA observed in PLCε+/+ mice was substantially attenuated in PLCε−/− mice. IL-1α is a member of the IL-1 family cytokines having pleiotropic functions, such as control of immune responses and inflammatory processes (30). On the other hand, TPA-induced up-regulation of mRNAs coding for the other molecules, including cyclooxygenase 2 (COX-2) and tumor necrosis factor-α (TNF-α), which are implicated in tumor promotion and inflammation (31, 32), was unaffected by the PLCε background.

Figure 3.

Role of PLCε in TPA-induced expression of proinflammatory molecules in vivo and in vitro. A, TPA-induced expression of representative proinflammatory molecules in the ear. Total cellular RNA was prepared from the ears of PLCε+/+ and PLCε−/− mice at the indicated time points after topical application of acetone vehicle (−) or TPA (+). The mRNA levels of the indicated proteins were semiquantitatively determined by RT-PCR. The amount of the template for each PCR was normalized with β-actin mRNA. B and C, TPA-induced expression of representative proinflammatory molecules in cultured skin cells. Dermal fibroblasts (B) and epidermal keratinocytes (C) from the dorsal skins of PLCε+/+ and PLCε−/− mice were stimulated with various concentrations of TPA for the indicated periods. The mRNA levels of the indicated proteins were semiquantitatively determined by RT-PCR. The amount of the template for each PCR was normalized with GAPDH or β-actin mRNA.

Figure 3.

Role of PLCε in TPA-induced expression of proinflammatory molecules in vivo and in vitro. A, TPA-induced expression of representative proinflammatory molecules in the ear. Total cellular RNA was prepared from the ears of PLCε+/+ and PLCε−/− mice at the indicated time points after topical application of acetone vehicle (−) or TPA (+). The mRNA levels of the indicated proteins were semiquantitatively determined by RT-PCR. The amount of the template for each PCR was normalized with β-actin mRNA. B and C, TPA-induced expression of representative proinflammatory molecules in cultured skin cells. Dermal fibroblasts (B) and epidermal keratinocytes (C) from the dorsal skins of PLCε+/+ and PLCε−/− mice were stimulated with various concentrations of TPA for the indicated periods. The mRNA levels of the indicated proteins were semiquantitatively determined by RT-PCR. The amount of the template for each PCR was normalized with GAPDH or β-actin mRNA.

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We next asked what kinds of cells in the skin are responsible for this effect. Because TPA is thought to diffuse through the epidermis to the dermis, keratinocytes, dermal fibroblasts, and resident leukocyte populations are candidate targets. Leukocytes became the first to be ruled out because PLCε was not detected in infiltrating Gr-1–positive granolocytes, B lymphocyte and non–B lymphocyte populations, or thioglycollate-elicited macrophages (Supplementary Fig. S1). This result suggested that keratinocytes and/or dermal fibroblasts are responsible for the role of PLCε in TPA-induced inflammation and led us to examine whether TPA-induced IL-1α up-regulation is recapitulated in keratinocytes and/or dermal fibroblasts in culture (Fig. 3B and C). In dermal fibroblasts, IL-1α mRNA expression was induced at 2 h after TPA application, and this induction was significantly compromised in fibroblasts from PLCε−/− mice (Fig. 3B). Expression of other representative proinflammatory molecules was not significantly affected by the PLCε background, except for macrophage inflammatory protein 2 (MIP-2), whose induction by higher concentration of TPA seemed to be attenuated in PLCε−/− fibroblasts. In sharp contrast, the IL-1α mRNA level in keratinocytes was not altered by TPA or the PLCε background (Fig. 3C). These results suggested that dermal fibroblasts, but not epidermal keratinocytes, play a major role in TPA-induced and PLCε-dependent up-regulation of IL-1α observed in the whole ears.

TPA induces activation of PLCε through Rap1 activation in dermal fibroblasts. Cultured dermal fibroblasts were used to analyze the molecular mechanism of TPA-dependent activation of PLCε. We first set up a method whereby the PLC activity is measured by increase in the cytosolic free calcium concentration triggered by the PLC product IP3. TPA application induced rapid increase in cytosolic free calcium, and this increase was totally abolished by pretreatment with a broad spectrum PLC inhibitor U73122 (Fig. 4A), indicating the complete dependency on PLC activity. When TPA-induced calcium increase was compared, PLCε−/− fibroblasts exhibited considerable reduction compared with PLCε+/+ fibroblasts, indicating that a major part of the calcium increase was accounted for by the activation of PLCε (Fig. 4B). In contrast, ATP-dependent calcium increase, which is mediated by PLCβ, was not affected by the PLCε background, confirming that no difference existed in the loading efficiency of Fura-2 AM between PLCε+/+ and PLCε−/− cells (Fig. 4B). Thus, TPA-induced cytosolic free calcium increase could be used as a good measure of PLCε activation.

Figure 4.

TPA-induced activation of PLCε in dermal fibroblasts. A, essential role of PLC in TPA-induced increase in the cytosolic free calcium concentration. After pretreatment with or without 2.5 μmol/L U73122 for 10 min, Fura-2–loaded dermal fibroblasts were stimulated with 100 ng/mL TPA at time point (*), followed with 10 μmol/L ionomycin at time point (**). Relative increases in the calcium concentrations were calculated as described in Materials and Methods. B, crucial role of PLCε in TPA-induced calcium increase. Fura-2–loaded dermal fibroblasts from PLCε+/+ and PLCε−/− mice were stimulated with 100 ng/mL TPA (top) or 250 μmol/L ATP (bottom) at time point (*), followed with 10 μmol/L ionomycin at time point (**).

Figure 4.

TPA-induced activation of PLCε in dermal fibroblasts. A, essential role of PLC in TPA-induced increase in the cytosolic free calcium concentration. After pretreatment with or without 2.5 μmol/L U73122 for 10 min, Fura-2–loaded dermal fibroblasts were stimulated with 100 ng/mL TPA at time point (*), followed with 10 μmol/L ionomycin at time point (**). Relative increases in the calcium concentrations were calculated as described in Materials and Methods. B, crucial role of PLCε in TPA-induced calcium increase. Fura-2–loaded dermal fibroblasts from PLCε+/+ and PLCε−/− mice were stimulated with 100 ng/mL TPA (top) or 250 μmol/L ATP (bottom) at time point (*), followed with 10 μmol/L ionomycin at time point (**).

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We next analyzed the upstream regulatory mechanism of TPA-induced PLCε activation. To this end, the activation statuses of Ras and Rap1, two major regulators of PLCε, were examined in TPA-stimulated fibroblasts by using the pull-down assays. As shown in Fig. 5A, TPA specifically induced activation of Rap1, but not Ras, regardless of the PLCε background. In contrast, EGF was capable of inducing activation of both Ras and Rap1. We next used siRNA-mediated knockdown of Rap1 expression to prove the role of Rap1 in TPA-dependent activation of PLCε. Because dermal fibroblasts expressed both Rap1A and Rap1B (Supplementary Fig. S2A), we transfected the cells with two different combinations of Rap1A-specific and Rap1B-specific siRNAs, both of which caused substantial reduction in the Rap1 expression (Supplementary Fig. S2B). As shown in Fig. 5B, transfection of the two siRNA combinations caused substantial reduction in TPA-induced cytosolic calcium increase and, hence, the activation of PLCε. The loading efficiency of Fura-2 AM was not affected by siRNA transfection, as no difference in the ATP-dependent calcium responses was shown. These results showed that Rap1 plays a crucial role in mediating the TPA-initiated signal, which leads to PLCε activation.

Figure 5.

Crucial role of Rap1 in TPA-induced PLCε activation. A, activation of Rap1 but not Ras by TPA stimulation. Dermal fibroblasts were stimulated with 100 ng/mL TPA or 20 ng/mL EGF for 5 min at room temperature. The levels of the GTP-bound forms of Ras and Rap1 were measured by the pull-down assays as described in Materials and Methods. One-tenth aliquots of the cell lysates used for the pull-down assays were subjected to Western blotting to show the equal inputs of Ras and Rap1. B, effects of Rap1 knockdown on TPA-induced PLCε activation. Dermal fibroblasts transfected with the indicated combinations of siRNAs were stimulated with 10 ng/mL TPA at time point (*), followed with 250 μmol/L ATP at time point (**) and finally with 10 μmol/L ionomycin at time point (***). Increase in the cytosolic free calcium was assayed as described in Fig. 4 as a measure of PLCε activation.

Figure 5.

Crucial role of Rap1 in TPA-induced PLCε activation. A, activation of Rap1 but not Ras by TPA stimulation. Dermal fibroblasts were stimulated with 100 ng/mL TPA or 20 ng/mL EGF for 5 min at room temperature. The levels of the GTP-bound forms of Ras and Rap1 were measured by the pull-down assays as described in Materials and Methods. One-tenth aliquots of the cell lysates used for the pull-down assays were subjected to Western blotting to show the equal inputs of Ras and Rap1. B, effects of Rap1 knockdown on TPA-induced PLCε activation. Dermal fibroblasts transfected with the indicated combinations of siRNAs were stimulated with 10 ng/mL TPA at time point (*), followed with 250 μmol/L ATP at time point (**) and finally with 10 μmol/L ionomycin at time point (***). Increase in the cytosolic free calcium was assayed as described in Fig. 4 as a measure of PLCε activation.

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TPA-induced Rap1 activation is mediated by RasGRP3 and PKC. To make a connection between the TPA-initiated signal and Rap1 activation, we focused on the DAG/TPA effector RasGRPs (15). Dermal fibroblasts express RasGRP1 and RasGRP3 (Supplementary Fig. S3A). Among them, RasGRP3 was the most likely candidate because RasGRP3, but not RasGRP1, is capable of activating Rap1 (16) and RasGRP2 is unresponsive to TPA (33). We therefore knocked down RasGRP3 expression by using siRNAs and examined Rap1 activity in TPA-stimulated cells. As expected, the knockdown of RasGRP3 expression by two different siRNAs substantially suppressed Rap1 activation when cells were stimulated with a low dose (1 ng/mL) of TPA (Fig. 6A). However, this inhibitory effect of RasGRP3 knockdown was overridden by a high dose (10 ng/mL) of TPA (Fig. 6B).

Figure 6.

Crucial roles of RasGRP3 and PKC in TPA-induced activation of Rap1. A, effect of RasGRP3-specific siRNAs on TPA-dependent activation of Rap1. Dermal fibroblasts were transfected with two different siRNAs (grp65 and grp67) targeting RasGRP3 or a negative control siRNA (Cont) and stimulated with or without 1 ng/mL TPA for 5 min. The Rap1-GTP levels were measured by the pull-down assay (top). Input Rap1 and actin levels in the cellular lysates were also shown. B, effect of a high concentration of TPA on the requirement of RasGRP3 for Rap1 activation. Dermal fibroblasts transfected with control siRNA or RasGRP3-specific siRNA (grp67) were stimulated with the indicated concentrations of TPA for 5 min. The cellular Rap1-GTP levels were measured by the pull-down assay. C, effect of a PKC inhibitor on TPA-induced activation of Rap1. After pretreatment with 5 μmol/L GF109203X (PKCi, +) or vehicle DMSO (−) for 15 min, dermal fibroblasts were stimulated with the indicated concentrations of TPA for 5 min, and the Rap1-GTP levels were determined by the pull-down assay. D, effect of a PKC inhibitor on TPA-induced activation of PLCε. After pretreatment as in C, the cells were stimulated with 100 ng/mL TPA at time point (*), followed with 10 μmol/L ionomycin at time point (**). The cytosolic free calcium concentrations were measured as described in Fig. 4.

Figure 6.

Crucial roles of RasGRP3 and PKC in TPA-induced activation of Rap1. A, effect of RasGRP3-specific siRNAs on TPA-dependent activation of Rap1. Dermal fibroblasts were transfected with two different siRNAs (grp65 and grp67) targeting RasGRP3 or a negative control siRNA (Cont) and stimulated with or without 1 ng/mL TPA for 5 min. The Rap1-GTP levels were measured by the pull-down assay (top). Input Rap1 and actin levels in the cellular lysates were also shown. B, effect of a high concentration of TPA on the requirement of RasGRP3 for Rap1 activation. Dermal fibroblasts transfected with control siRNA or RasGRP3-specific siRNA (grp67) were stimulated with the indicated concentrations of TPA for 5 min. The cellular Rap1-GTP levels were measured by the pull-down assay. C, effect of a PKC inhibitor on TPA-induced activation of Rap1. After pretreatment with 5 μmol/L GF109203X (PKCi, +) or vehicle DMSO (−) for 15 min, dermal fibroblasts were stimulated with the indicated concentrations of TPA for 5 min, and the Rap1-GTP levels were determined by the pull-down assay. D, effect of a PKC inhibitor on TPA-induced activation of PLCε. After pretreatment as in C, the cells were stimulated with 100 ng/mL TPA at time point (*), followed with 10 μmol/L ionomycin at time point (**). The cytosolic free calcium concentrations were measured as described in Fig. 4.

Close modal

We next examined the role of PKCs on TPA-induced Rap1 activation. One reason for doing this was because it had been reported that RasGRP3 needs phosphorylation by certain isozymes of PKCs for its GEF activity (15, 16, 34). As shown in Fig. 6C and D, pretreatment with the PKC inhibitor GF109203X potently suppressed not only the TPA-dependent Rap1 activation but also the TPA-induced PLCε activation. This inhibition could not be overridden by stimulation with high doses of TPA. These results indicated that the TPA-initiated signal for Rap1 activation is mediated by both RasGRP3 and PKCs.

The causal relationship between tumor promotion and inflammation has been supported from the observations that tumor promotion can result from exposure of initiated cells to chemical irritant, such as phorbol esters, chronic irritations, and inflammation and that phorbol esters are strong inducers of inflammatory reactions (23). This old theme has gained experimental supports at the molecular level from recent studies using genetically engineered mice. For instance, mice lacking TNF-α, a potent proinflammatory cytokine, are resistant to skin carcinogenesis by the DMBA/TPA protocol (32). Inhibition of inflammation by pharmacologic or genetic inactivation of COX-2 reduced tumor formation and malignant progression in mice carrying the mutated Apc gene (31). The importance of innate immunity was also shown by reduced tumor formation in the MyD88-deficient mice (35, 36). However, intracellular signaling mechanisms mediating this relationship remain largely unsolved.

In this study, we have shown that PLCε−/− mice exhibit substantially attenuated inflammatory responses to TPA treatment in the skin. Considering that the same mice exhibited marked resistance to tumor formation in the two-stage skin carcinogenesis and to TPA-induced basal layer cell proliferation and epidermal hyperplasia (14), our results suggest that PLCε may be involved in an intracellular signaling pathway linking tumor promotion and inflammation. Because cultured keratinocytes and dermal fibroblasts, which are free from inflammatory cells, exhibited no difference in proliferation potential depending on the PLCε background, infiltrating inflammatory cells, whose number is reduced in PLCε−/− mice, are likely to be the main source of cytokines and growth factors, which are shown to be directly or indirectly, via stimulation of stromal cells, involved in the TPA-induced keratinocyte proliferation and possibly in tumor promotion (37). Thus, PLCε is likely to be involved in an initial phase of inflammation, i.e. recruitment of inflammatory leukocytes to the site of TPA administration. Because PLCε is found to be not expressed in leukocytes, including granulocytes, macrophages, and lymphocytes, keratinocytes and/or dermal fibroblasts are the possible sources of factors involved in this process. Search for such candidate proinflammatory molecules has identified IL-1α, whose induction by TPA is compromised in the skin of PLCε−/− mice. Experiments using cell culture have shown that dermal fibroblasts may be the main source of IL-1α induced by TPA in a PLCε-dependent manner. IL-1α, as well as IL-1β, is a member of the IL-1 cytokine family having pleiotropic functions (30). IL-1α exerts its activity by binding to its cell surface receptors belonging to the Toll-like receptor/IL-1 receptor superfamily (38). Binding of IL-1α to its receptors induces recruitment of MyD88, TRAF6, and protein kinases, including IRAK and IKK, leading to activation of the nuclear factor-κB (NF-κB)–dependent transcription of proinflammatory cytokines, including IL-6, KC, MIP-2, TNFα, and IL-1 (38). IL-1α also up-regulates expression of cell adhesion molecules, such as VCAM-1 and E-selectin on endothelial cells and ICAM-1 on mesenchymal cells, which are required for inflammatory responses, including transmigration of leukocytes to the sites of inflammation (30). Furthermore, it was reported that both epidermal hyperplasia induced by TPA and tumor promotion by repeated treatment with DMBA are suppressed by injection of anti–IL-1α antibody, indicating a crucial role of IL-1α (39, 40). Thus, it is likely that PLCε plays a crucial role in TPA-induced up-regulation of IL-1α in dermal fibroblasts, which initiates a series of inflammatory reactions in the skin. Also, our result has raised an interesting possibility that the microenvironment facilitating proliferation of the initiated cells may be provided by their surrounding cells through induction of inflammation mediated by PLCε. Our preliminary study showed that intestinal tumor formation in ApcMin mice was suppressed on the PLCε−/− background,1

1

M. Li, H. Edamatsu, and T. Kataoka, unpublished data.

suggesting rather universal role of PLCε in tumorigenesis.

We have revealed that Rap1 plays a crucial role in mediating TPA-dependent PLCε activation in dermal fibroblasts. Rap1 activation by TPA seems to be mediated by RasGRP3, which is a DAG/TPA-regulated Rap1-GEF expressed in dermal fibroblasts. Contrary to the previous reports that RasGRP3 is capable of activating Ras, as well as Rap1 (15, 16), we have failed to detect activation of Ras by TPA in our system; the reason for which is currently unknown. Furthermore, PKCs are also found to be required for TPA-dependent activation of Rap1 and PLCε. It has been reported that certain isozymes of PKCs phosphorylate RasGRP3 at its Thr133, and this phosphorylation is required for the GEF activity toward Ras (15, 16). Although a similar mechanism has not been proved with Rap1 as substrate, it may well be involved in regulation of the Rap1-GEF activity of RasGRP3. This is supported from a three-dimensional model structure of the GEF domain of RasGRP3, which implies that the phosphorylation of Thr133 plays an important role in a conformational change enhancing its guanine nucleotide exchange activity (34). Thus, we speculate that TPA may induce Rap1 activation in two ways: through direct activation of RasGRP3 and through direct activation of PKCs, which in turn phosphorylates and activates RasGRP3. We have observed that stimulation with a high dose (10 ng/mL) of TPA overrides the inhibitory effect of siRNAs against RasGRP3 but not of the PKC inhibitor on Rap1 activation. This phenomenon may be accounted for by postulating that RasGRP3 activity is fully dependent on PKC phosphorylation and that a high dose of TPA induces substantial activation of the residual RasGRP, which survived the siRNA action, through PKC activation so that the RasGRP3 activity reaches saturation. However, we cannot rule out the possibility that other PKC-regulated mechanisms of Rap1 activation may be involved in this process. Because the actual concentration of TPA reaching the dermis is totally unknown, physiologic implication of these results is very difficult. The transcription of the IL-1α gene is under very complex regulation involving not only NF-κB (41) and activator protein-1 (42) but also DNA methylation (43). Although NF-κB seems to be the most likely molecule involved in the TPA-induced and PLCε-mediated IL-1α expression, we have observed no alteration in the TPA-dependent induction of other NF-κB–regulated genes, such as IL-6, TNF-α (38), and COX-2 (44). Thus, the signaling mechanism downstream of PLCε leading to the IL-1α expression needs further clarification.

Our results raise the possibility that RasGRP3 may also play a crucial role in tumor promotion by TPA. However, the role of RasGRP3 in tumor promotion has not been studied, although mice whose RasGRP3 gene was disrupted were already generated (45). In dermal fibroblasts, we have detected α, δ, and ε isozymes of PKC by Western blotting.2

2

S. Ikuta, H. Edamatsu, M. Li, L. Hu, and T. Kataoka, unpublished data.

However, α and δ isozymes are thought to be antitumorigenic (1821), and the role of ε isozyme in dermal fibroblasts has not been well documented. Further studies on the role of various PKC isozymes in Rap1-PLCε signaling will be required for elucidation of the molecular mechanism linking tumor promotion and inflammation by TPA.

Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

S. Ikuta and H. Edamatsu contributed equally to this work.

Grant support: Grant-in-aids for Priority Areas 17014061 (T. Kataoka) and for Scientific Research 17390078 (T. Kataoka) and 18790223 (H. Edamatsu), 21st Century COE and Global COE Programs (T. Kataoka), and grant for Initiatives for Attractive Education in Graduate Schools (M. Li) from the Ministry of Education, Science, Sports and Culture of Japan; grant for the Program for Promotion of Fundamental Studies of Health Sciences 06-3 (T. Kataoka) from the National Institute of Biomedical Innovation; and grant from Hyogo Science and Technology Association (H. Edamatsu).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Dr. Masahiro Oka for helpful discussions and Tadashi Murase for setting up the assay system for cytosolic calcium concentration.

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Supplementary data