Keratinocyte proliferation

Keratinocyte proliferation DEFAULT

Regulation of keratinocyte proliferation

1. In physiological situations the proliferation of epidermal cells (keratinocytes) in the skin is a tightly controlled process. 2. However, in many common skin diseases, such as in psoriasis, the control mechanisms go awry resulting in pathological epidermal hyperplasia (thickening). 3. In those situations the keratinocytes enter the alternative pathway of proliferation characterized by excessive growth rate, aberrant responses to growth factors, faulty differentiation, and increased migratory capacity. 4. The participation of different growth factors in enhancing or inhibiting keratinocyte growth, both in physiological and pathological conditions, has been reviewed. 5. The regulatory processes governing epidermal growth have relevance for the understanding of the mechanism of action of the drugs used in the treatment of skin diseases associated with epidermal hyperplasia.

Sours: https://pubmed.ncbi.nlm.nih.gov/9559309/

Abstract

The biological effects of 1,25-dihydroxyvitamin D3 are mediated by a nuclear receptor, the vitamin D receptor (VDR). Targeted ablation of the VDR in mice results in hypocalcemia, hypophosphatemia, hyperparathyroidism, rickets, osteomalacia, and alopecia. Normalization of mineral ion homeostasis prevents these abnormalities with the exception of the alopecia. Because 1,25(OH)2D3 has been shown to play a role in keratinocyte proliferation and differentiation, we undertook studies in primary keratinocytes and skin isolated from VDR null mice to determine if a keratinocyte abnormality could explain the alopecia observed. The basal proliferation rate of the VDR null and wild-type keratinocytes was identical both under proliferating and differentiating conditions. Assessment of in vivo keratinocyte proliferation at 4 days of age confirmed that VDR ablation did not have a significant effect. There was no difference in the basal expression of markers of keratinocyte differentiation (keratin 1, involucrin, and loricrin) in the keratinocytes isolated from VDR-ablated mice when compared with those isolated from control littermates. Similarly, in vivo expression of these genes was not altered at 4 days of age. When anagen was induced by depilation at 18 days of age, the VDR null mice had a profound impairment in initiation of the hair cycle. These data suggest that the alopecia in the VDR null mice is not attributable to an intrinsic defect in keratinocyte proliferation or differentiation, but rather to an abnormality in initiation of the hair cycle.

THE BIOLOGICALLY active metabolite of vitamin D, 1,25-dihydroxyvitamin D (1,25(OH)2D3), interacts with target genes by binding to a nuclear receptor, the vitamin D receptor (VDR) (reviewed in Ref. 1). Targeted ablation of the DNA-binding domain of the VDR in mice results in hypocalcemia, hypophosphatemia, hyperparathyroidism, rickets, osteomalacia, and alopecia (2, 3). Normalization of mineral ion homeostasis by a diet high in lactose, calcium and phosphorus, normalizes this phenotype with the exception of alopecia (2). Alopecia is not a feature of profound dietary vitamin D deficiency, nor is it observed in kindreds with 25-hydroxyvitamin D3 1α-hydroxylase mutations. The development of alopecia in mice and humans with VDR mutations but not with ligand deficiency remains unexplained.

1,25(OH)2D3 has been shown to be a potent inhibitor of keratinocyte proliferation. 1,25(OH)2D3 also stimulates keratinocyte differentiation in a concentration-dependent manner as evidenced by enhanced formation of cornified envelopes and induction of marker gene expression (4, 5).

The hair follicle consists of mesenchymal cells and keratinocytes that form the bulb region, deep in the hypodermal fat (6). The outermost keratinocytes give rise to the outer root sheath, the inner root sheath and the hair shaft. Mice are born without hair although histological examination of the skin at the time of birth reveals immature hair follicles. Numerous anagen follicles are evident within the first 6 days of life and by 15 days the hair cycle enters the catagen phase, progressing to the telogen phase. Subsequently, the follicle undergoes cycles of growth (anagen), regression (catagen), and rest (telogen). In vivo, VDR expression in the outer root sheath keratinocytes correlates with decreased keratinocyte proliferation and increased differentiation in late anagen and catagen (7). We hypothesized that ablation of the VDR might alter the proliferation and differentiation of keratinocytes, and thereby lead to the alopecia observed in humans and mice with VDR mutations. Primary keratinocytes were, therefore, isolated from VDR null mice and control littermates to examine their proliferation rate and expression of keratinocyte differentiation markers. We also examined the expression of two candidate genes which, when misexpressed, lead to alopecia: PTH-related protein (PTH-rP) and hairless (hr).

PTH-rP, initially discovered as the cause of humoral hypercalcemia of malignancy (8), is expressed in a wide variety of normal cells, including epidermal keratinocytes and has been implicated in keratinocyte differentiation (9). Forced overexpression of PTH-rP in keratinocytes of normal mice interferes with normal hair follicle development (10). Consistent with these observations, bPTH (7–34), an antagonist of the PTH/PTH-rP receptor, has been shown to increase the number and length of hair shafts in SKH-1 hairless mice (11). Because the expression of PTH-rP in human keratinocytes is suppressed by 1,25(OH)2D3, it is possible that overexpression of this hormone by the keratinocytes of the VDR knockout mice contributes to the alopecia observed.

The second potential candidate gene we examined was the hairless gene. Like the VDR knockout mice, hairless (hr/hr) mice have a normal first hair coat, however, they develop alopecia at approximately 2 weeks of age (12). Furthermore, like the VDR knockout mice, the alopecia is accompanied by the presence of large dermal cysts. Mutations of this gene in humans have been shown to be the cause of congenital atrichia in several families (13).

Although many genes implicated in hair follicle development are expressed in the mature hair follicle, it is thought that factors that control folliculogenesis are distinct from the factors responsible for the regulation of the hair cycle. The first coat of hair is dependent on factors that control development, whereas subsequent coats are dependent on normal cycling of hair follicles. Therefore, we also performed studies to examine whether the alopecia in the VDR-ablated mice was secondary to a hair cycle defect.

Materials and Methods

Animal maintenance

All studies performed were approved by the institutional animal care committee. VDR null mice and control littermates were maintained in a virus- and parasite-free barrier facility and exposed to 12-h light, 12-h dark cycle. The heterozygous mothers were fed autoclaved Purina rodent chow (5010, Ralston Purina Co., St. Louis, MO) containing 1% calcium, 0.67% phosphorus, 0% lactose, and 4.4 IU vitamin D/g (regular diet). Upon weaning at 18 days of age, VDR null mice and control littermates were fed a γ-irradiated test diet (TD96348, Teklad, Madison, WI, containing 2% calcium, 1.25% phosphorus, and 20% lactose supplemented with 2.2 IU vitamin D/g) which has been shown to prevent abnormalities in mineral ion homeostasis in VDR-ablated mice (2).

Cell culture

Primary keratinocytes were isolated from 2- to 3-day-old receptor-ablated mice and control littermates by a trypsin floating procedure as previously described (14). Briefly, the skin was isolated and floated on 0.25% trypsin (Life Technologies, Inc., Grand Island, NY) at 4 C overnight. The epidermis was then separated from the dermis, minced with scissors and stirred in MEM with 4% Chelex-treated FCS (HyClone Laboratories, Inc., Logan, UT), epidermal growth factor (EGF; 10 ng/ml; Collaborative Research, Inc., Cambridge, MA) and 0.05 mm CaCl2 (low calcium medium) for 1 h at 4 C. The cell suspension derived from two mice of the same genotype was filtered through three layers of gauze and plated in low calcium medium in collagen (Vitrogen 100, Palo Alto, CA)-coated 100-mm dishes and incubated at 34 C, 8% CO2. After achieving 80% confluence, these cells were reseeded at 2.5 × 105 cells/well of a 6-well plate in low calcium medium and grown to 80% confluence before addition of 1,25(OH)2D3 (10−8m) and/or inducing differentiation by increasing the calcium concentration to 2.0 mm. Forty hours later, total RNA was prepared for northern analysis.

[3H]thymidine incorporation

Keratinocytes were plated at 5.0 × 104 cells/well of a 24-well plate and grown to 80% confluence in keratinocyte growth medium containing 0.05 mm calcium. Cells were then treated with high calcium and/or 10−8m 1,25(OH)2D3. After 24 h, cells were labeled for 12 h with[ 3H]thymidine (NEN Life Science Products, Boston, MA) and radionucleotide incorporation was assessed as previously described (15). The[ 3H]thymidine counts/min (cpm) were corrected for protein concentration.

BrdU incorporation

Mice were injected ip with 5-bromo-2′-deoxyuridine (Brdu) (250 mg/kg; Sigma, St. Louis, MO) and 5-fluoro-2′-deoxyuridine (Fdu) (30 mg/kg; Sigma). Animals were killed 2 h after Brdu/Fdu injection. Skin specimens were obtained from the middorsum of VDR-ablated mice and control littermates. After fixation for 3 h in 4% formaldehyde in PBS (pH 7.2), specimens were processed, embedded in paraffin, and cut into 6-μm sections with a Leica Corp. RM 2025 microtome (Leica Corp., Deerfield, IL). Brdu staining was performed using a Brdu Staining Kit (Zymed Laboratories, Inc., South San Francisco, CA) following the manufacturer’s instructions.

RNA isolation and Northern blot analysis

Total RNA was isolated from cultured keratinocytes and mouse skin using TRI Reagent (Sigma) according to the manufacturer’s instructions. For Northern analysis, total RNA (3–20μ g) was electrophoresed through 1% agarose-formaldehyde gels and transferred to Biotrans nylon membranes (ICN Pharmaceuticals, Inc., Irvine, CA) in 20 × SSC. The membranes were hybridized with complementary DNA (cDNA) probes labeled with[α -32P]dATP (NEN Life Science Products) or with a γ-32P-labeled antisense oligonucleotide probe for 18S ribosomal RNA. The cDNA probes used were a 0.7- kb EcoRI-XhoI fragment of mouse keratin-1, a 2.1-kb EcoRI-XhoI fragment of mouse involucrin, a 1.4-kb EcoRI-XhoI fragment of mouse loricrin (from American Type Culture Collection, Manassas, VA), a 0.4- kb AvrII-SmaI fragment of mouse PTH-rP and a 0.9-kb BamHI-HindIII fragment of mouse hairless (gift from Dr. Jonathan P. Stoye, National Institute for Medical Research, Mill Hill, London, NW71AA, UK). Hybridization was carried out at 68 C in QuikHyb (Stratagene, La Jolla, CA). These probes all generated single transcripts of the appropriate size. The intensity of the messenger RNA (mRNA) bands was assessed by a Cyclone Storage Phosphor System (Packard Instrument Company, Meriden, CT) using Opti Quant software. 18S ribosomal RNA signals were used to normalize for variations in RNA loading.

Anagen induction

Under avertin-induced general anesthesia, 18-day-old receptor- ablated mice and control littermates were subjected to depilation of their dorsal hair using Wax Strips (Del Laboratories, Farmingdale, NY) following the manufacturer’s instructions. Histological examination was performed 6 and 10 days following this procedure.

Statistical analyses

Data are presented as the mean ± sem. Student’s unpaired t test was used to identify significant differences (P < 0.05).

Results

Primary keratinocytes maintained in low calcium (0.05 mm), behave like cells of the basal layers of the epidermis. An increase in the extracellular calcium concentration of cultured keratinocytes triggers growth arrest and induces a program of terminal differentiation similar to that observed in the suprabasal layers of the epidermis (14). When cultured in either low or high calcium medium, the keratinocytes from the VDR-ablated mice maintained the same proliferative rate as those of wild-type control littermates (Fig. 1A).

Figure 1.

Role of the VDR in keratinocyte proliferation. A, Proliferation of VDR+/+ and VDR−/− keratinocytes. Second passage keratinocytes were grown to 80% confluence in keratinocyte growth medium containing 0.05 mm calcium.[ 3H]thymidine incorporation (corrected for protein concentration) was assessed in low calcium media and 24 h after high calcium (2.0 mm) and/or 10−8m 1,25(OH)2D3 treatment. Data represent the means of triplicate wells ± sem from at least three independent experiments. WT, Wild-type; HOM, homozygous. B, Brdu incorporation into hair bulb keratinocytes. Skin sections were obtained from the mid dorsum of 4-day-old littermates 2 h post ip injection of Brdu. Following immunostaining with an anti-Brdu antibody, Brdu incorporation was assessed in the keratinocytes of the hair follicle bulbs at the level of the dermal papilla. Data represents the mean ± sem of the percentage of Brdu postive cells in 30 hair follicle bulbs from each of 4 homozygous and wild-type littermates. WT, Wild-type; HOM, homozygous.

Figure 1.

Role of the VDR in keratinocyte proliferation. A, Proliferation of VDR+/+ and VDR−/− keratinocytes. Second passage keratinocytes were grown to 80% confluence in keratinocyte growth medium containing 0.05 mm calcium.[ 3H]thymidine incorporation (corrected for protein concentration) was assessed in low calcium media and 24 h after high calcium (2.0 mm) and/or 10−8m 1,25(OH)2D3 treatment. Data represent the means of triplicate wells ± sem from at least three independent experiments. WT, Wild-type; HOM, homozygous. B, Brdu incorporation into hair bulb keratinocytes. Skin sections were obtained from the mid dorsum of 4-day-old littermates 2 h post ip injection of Brdu. Following immunostaining with an anti-Brdu antibody, Brdu incorporation was assessed in the keratinocytes of the hair follicle bulbs at the level of the dermal papilla. Data represents the mean ± sem of the percentage of Brdu postive cells in 30 hair follicle bulbs from each of 4 homozygous and wild-type littermates. WT, Wild-type; HOM, homozygous.

Like calcium, 1,25(OH)2D3 has also been shown to decrease proliferation and induce keratinocyte differentiation (4, 14, 16). Although 10−8m 1,25(OH)2D3 suppressed the proliferation rate of wild-type keratinocytes maintained in low calcium (68 ± 3.6% of control), no modulation of proliferation was observed in the VDR null keratinocytes. When an increase in extracellular calcium was added to induce terminal differentiation and growth arrest, [3H] thymidine incorporation rate decreased and 10−8m 1,25(OH)2D3 had no effect on proliferation. These data suggest that the absence of a functional VDR does not effect the proliferation rate of terminally differentiated keratinocytes. To confirm that this is also the case in vivo, proliferation of hair follicle keratinocytes was assessed by Brdu incorporation at 4 days of age, correlating with the first anagen. There was no genotype-dependent difference in the number of BrDu labeled keratinocytes in the hair bulb (Fig. 1B), follicle, or interfollicular region (data not shown). These data confirmed the in vitro findings that the proliferation rate of keratinocytes lacking functional VDRs did not differ significantly from that of wild-type keratinocytes.

The expression of differentiation markers in VDR null and wild-type keratinocytes was then examined. Keratin 1 (K1) is an early marker of differentiation that is expressed in the spinous layer of the epidermis. Involucrin, a component of the cornified envelope, is a suprabasal marker of keratinocyte differentiation that is expressed in the late spinous layer and throughout the granular layer. Loricrin is a marker of the granular and cornified layers (17). All three differentiation markers were expressed equally in the VDR null and wild-type keratinocytes under both proliferative (0.05 mm CaCl2) and differentiating (2 mm CaCl2) conditions (Fig. 2A). Although the proliferation rate of the keratinocytes decreased with addition of high CaCl2 media, the only keratinocyte differentiation marker that changed significantly was involucrin. Although the circulating levels of 1,25-dihydroxyvitamin D in suckling mice are not suppressed (3; and data not shown), it is possible that high intracellular levels may be present in keratinocytes, which may lead to differential expression of hormonally regulated genes in vivo. To ascertain that the observations in primary keratinocytes reflected the in vivo state, we examined expression of these markers in skin mRNA isolated from VDR-ablated mice and wild-type littermates. Northern analyses revealed similar levels of expression of keratin 1, involucrin and loricrin at 4 days of age (Fig. 2B).

Figure 2.

Expression of markers of keratinocyte differentiation. A, Keratinocytes were grown to 80% confluence in 0.05 mm calcium, then treated or not with 2.0 mm calcium. Cells were harvested after 40 h. Three micrograms of total RNA was used for Northern analysis. Data represents the mean± sem of at least four independent experiments performed with keratinocytes isolated independently from at least three mice of each genotype. Data are normalized to wild-type low calcium, and in each case the intensity of the signal is corrected for that of the 18S rRNA. WT, Wild-type; HOM, homozygous. B, Total RNA was isolated from the skin of 4-day-old littermates. Northern analysis was performed using 15 μg of skin RNA. Membranes were sequentially hybridized with mouse keratin 1, involucrin and loricrin cDNA probes. 18S ribosomal RNA signals were used to normalize for RNA loading. Data are expressed as percentage of wild-type control and represents the mean ± sem of Northern analyses performed with skin RNA isolated from four mice of each genotype. WT, Wild-type; HOM, homozygous.

Figure 2.

Expression of markers of keratinocyte differentiation. A, Keratinocytes were grown to 80% confluence in 0.05 mm calcium, then treated or not with 2.0 mm calcium. Cells were harvested after 40 h. Three micrograms of total RNA was used for Northern analysis. Data represents the mean± sem of at least four independent experiments performed with keratinocytes isolated independently from at least three mice of each genotype. Data are normalized to wild-type low calcium, and in each case the intensity of the signal is corrected for that of the 18S rRNA. WT, Wild-type; HOM, homozygous. B, Total RNA was isolated from the skin of 4-day-old littermates. Northern analysis was performed using 15 μg of skin RNA. Membranes were sequentially hybridized with mouse keratin 1, involucrin and loricrin cDNA probes. 18S ribosomal RNA signals were used to normalize for RNA loading. Data are expressed as percentage of wild-type control and represents the mean ± sem of Northern analyses performed with skin RNA isolated from four mice of each genotype. WT, Wild-type; HOM, homozygous.

The effects of 1,25(OH)2D3 on the expression of these markers was then examined in cultured keratinocytes. Treatment with 10−8m 1,25(OH)2D3 did not affect the expression of keratin 1 and loricrin. However, the expression of involucrin in wild-type keratinocytes, but not receptor-ablated cells, was markedly decreased by 1,25(OH)2D3 under proliferating (0.05 mm CaCl2, not shown) and differentiating (2.0 mm CaCl2) conditions, respectively (Fig. 3A). The suppression of involucrin expression by 1,25(OH)2D3 is a novel observation and may be a species-specific phenomenon. Previous studies in human keratinocytes have demonstrated induction of this gene by 1,25(OH)2D3, correlating with the differentiation promoting effects of this steroid hormone (4).

Figure 3.

Effect of 1,25(OH)2D3 on involucrin (A) and PTH-rP (B) mRNA expression. Second passage keratinocytes were grown to 80% confluence before increasing the calcium concentration to 2.0 mm and/or adding 1,25(OH)2D3 (10−8m). Forty hours later RNA was isolated. Ten micrograms of total RNA was used for Northern analysis. The membrane was hybridized with mouse involucrin cDNA or mouse PTH-rP probes. 18S ribosomal RNA signals were used to normalize for RNA loading. Data represents the mean ± sem of three independent experiments performed with keratinocytes isolated independently from three mice of each genotype. One representative Northern is shown for each probe, along with the 18S ribosomal RNA autoradiogram. WT, Wild-type; HOM, homozygous.

Figure 3.

Effect of 1,25(OH)2D3 on involucrin (A) and PTH-rP (B) mRNA expression. Second passage keratinocytes were grown to 80% confluence before increasing the calcium concentration to 2.0 mm and/or adding 1,25(OH)2D3 (10−8m). Forty hours later RNA was isolated. Ten micrograms of total RNA was used for Northern analysis. The membrane was hybridized with mouse involucrin cDNA or mouse PTH-rP probes. 18S ribosomal RNA signals were used to normalize for RNA loading. Data represents the mean ± sem of three independent experiments performed with keratinocytes isolated independently from three mice of each genotype. One representative Northern is shown for each probe, along with the 18S ribosomal RNA autoradiogram. WT, Wild-type; HOM, homozygous.

PTH-rP is expressed in epidermal keratinocytes and has been implicated in the regulation of hair growth (9, 11). Because the expression of PTH-rP in human keratinocytes is suppressed by 1,25(OH)2D3 (18) it is possible that overexpression of this hormone by the keratinocytes of the VDR knockout mice could lead to alopecia. To clarify this hypothesis, we examined the expression of PTH-rP in cultured keratinocytes isolated from VDR null mice and wild-type littermates. PTH-rP was expressed at normal levels in the VDR null cells (Fig. 3B). The expression of PTH-rP in wild-type keratinocytes, but not receptor-ablated cells, was suppressed by 10−8m 1,25(OH)2D3 under proliferating (0.05 mm CaCl2 not shown) and differentiating (2.0 mm CaCl2) conditions, respectively (Fig. 3B). Therefore, although repression of PTH-rP mRNA by 1,25(OH)2D3 is VDR-dependent, the basal expression of this hormone is not altered in VDR null keratinocytes.

Mutation of Hr (hairless) has been shown to cause alopecia in both mice (12) and humans (13). Like the VDR knockout mice, hairless (hr/hr) mice have a normal first coat of hair then develop alopecia accompanied by the presence of large dermal cysts. It is possible, therefore, that reduced expression of this gene by the keratinocytes of the VDR null mice could be responsible for the alopecia observed. However, the expression of hr was found to be normal in both cultured keratinocytes and skin isolated from VDR null mice (Fig. 4).

Figure 4.

Expression of the hairless gene. A, Keratinocytes were grown to 80% confluence in 0.05 mm calcium, then treated (or not) with 2.0 mm calcium. Cells were harvested after 40 h. Twenty micrograms of total RNA was used for Northern analysis. Data represents the mean ± sem of three independent experiments performed with keratinocytes isolated independently from three mice of each genotype. Data are normalized to wild-type low calcium and in each case the intensity of the signal is corrected for that of the 18S rRNA. WT, Wild-type; HOM, homozygous. B, Total RNA was isolated from the skin of 4-day-old littermates. Northern analysis was performed using 15 μg of skin RNA. 18S ribosomal RNA signals were used to normalize for RNA loading. Data represents the mean ± sem of Northern analyses performed with skin RNA isolated from four mice of each genotype. WT, Wild-type; HOM, homozygous.

Figure 4.

Expression of the hairless gene. A, Keratinocytes were grown to 80% confluence in 0.05 mm calcium, then treated (or not) with 2.0 mm calcium. Cells were harvested after 40 h. Twenty micrograms of total RNA was used for Northern analysis. Data represents the mean ± sem of three independent experiments performed with keratinocytes isolated independently from three mice of each genotype. Data are normalized to wild-type low calcium and in each case the intensity of the signal is corrected for that of the 18S rRNA. WT, Wild-type; HOM, homozygous. B, Total RNA was isolated from the skin of 4-day-old littermates. Northern analysis was performed using 15 μg of skin RNA. 18S ribosomal RNA signals were used to normalize for RNA loading. Data represents the mean ± sem of Northern analyses performed with skin RNA isolated from four mice of each genotype. WT, Wild-type; HOM, homozygous.

These data suggest that the VDR-ablated keratinocytes possess the same proliferative and differentiation potential as wild-type cells. Because the factors that control follicle morphogenesis in utero and the initial hair coat are different from factors that regulate hair cycling, we tested the response of the VDR knockout mice to anagen induction by depilation at 18 days of age, a time when there is no histological difference in the skin of the wild-type and VDR null mice. Because these animals are in the C57BL-6 background, this procedure should result in progressive skin pigmentation and thickening with 5 to 6 days, correlating with induction of anagen. Mature anagen follicles should be present within 10 days, accompanied by the appearance of hair shafts. Brdu incorporation performed 6 days post anagen induction (24 days of age), revealed marked Brdu incorporation in the follicle keratinocytes of the wild-type control littermates, correlating with marked proliferation that characterizes anagen (Fig. 5A). Control sections obtained from skin not subjected to depilation did not reveal induction of anagen follicles or skin thickening (data not shown). In contrast, rare Brdu positive cells were seen among the hair follicle keratinocytes of the VDR null mice (Fig. 5B). Also notable at this stage is the absence of skin thickening and lack of anagen follicles in the receptor-ablated animals. Even 10 days post depilation, a time when hair shafts are being formed in the wild-type littermates (Fig. 5C, arrows), there was no clinical or histological evidence of anagen induction in the VDR null mice (Fig. 5D). This data suggests that the alopecia in the VDR null mice is secondary to a defect in initiation of the hair cycle. Although clinically, alopecia totalis is not observed in the receptor-ablated mice until approximately 100 days of age, these studies demonstrate that the defect is present at 18 days of age. It is of note that the wild-type littermates have no evidence of having been subjected to depilation after 3 weeks, whereas the VDR-ablated mice remain profoundly alopecic 13 weeks post depilation.

Figure 5.

Response of VDR null mice to anagen induction. Littermates, fed a diet which maintains normal mineral ion homeostasis, were subjected to depilation at 18 days of age. Brdu incorporation into follicle keratinocytes was evaluated 6 days later in wild-type (A) and receptor-ablated littermates (B). Routine histology, was performed 10 days later (C, wild-type; D, homozygous-ablated mice). Data are representative of those obtained using three mice of each genotype for each time point. Magnification 10×.

Figure 5.

Response of VDR null mice to anagen induction. Littermates, fed a diet which maintains normal mineral ion homeostasis, were subjected to depilation at 18 days of age. Brdu incorporation into follicle keratinocytes was evaluated 6 days later in wild-type (A) and receptor-ablated littermates (B). Routine histology, was performed 10 days later (C, wild-type; D, homozygous-ablated mice). Data are representative of those obtained using three mice of each genotype for each time point. Magnification 10×.

Discussion

The observation that the histological appearance of the skin cannot reliably predict the presence of keratinocyte defects (19), combined with the previously reported effects of 1,25 on keratinocyte proliferation and differentiation (4), suggested that the alopecia in the VDR null mice may be secondary to a keratinocyte defect. Our studies, however, suggest that keratinocytes lacking functional VDRs possess the same proliferative and differentiation potential as wild-type cells. These studies, however, cannot rule out a defect in the VDR deficient keratinocytes related to their role in stimulating dermal papilla cells or receiving signals from these cell, required for the maintenance of the normal hair cycle. We, therefore, examined the response of the VDR null mice to initiation of anagen. These studies revealed that the VDR null mice had failure of anagen initiation, and remained alopecic postwaxing. Therefore, although hair follicle development is normal in the absence of a functional VDR, hair cycling is abnormal secondary to defect in anagen initiation. The cellular and molecular basis for this an abnormality is currently unclear.

Hair follicle development during embryogenesis requires a series of reciprocal interactions between the epithelium and the underlying mesenchymal cells. Initially, the dermal mesenchyme signals the epithelium to form the epidermal placode. The epithelium then sends a message to the underlying mesenchyme to initiate mesenchymal condensation. In response to signals from the condensed mesenchyme, hair elongation is observed. Recent studies have identified fibroblast growth factors, bone morphogenic proteins, and sonic hedgehog as epithelial-derived signaling molecules in the early stages of hair follicle morphogenesis. Postnatally, the maintenance of normal hair is dependent on the integrity of the dermis, epidermis, and normal hair cycles. Each hair follicle perpetually goes through three stages: growth (anagen), involution (catagen), and rest (telogen). Normal cycling of hair follicles is dependent on the interaction of the follicular keratinocytes with the mesenchymal dermal papilla cells. At the initiation of anagen, signals, thought to originate from the dermal papilla cells, induce the epithelial cells of the hair follicle to proliferate resulting in the full-length anagen follicle. These cells then differentiate to form the mature hair follicle, which includes the outer root sheath, the inner root sheath, and the hair shaft. The anagen follicle subsequently receives a signal that results in the initiation of catagen, characterized by apoptosis of the lower part of the hair follicle (20). The follicle then goes through the telogen, or resting, phase until the initiation of the following anagen by factors thought to emanate from the dermal papilla cells.

The protein products of several genes important for hair morphogenesis, such as insulin-like growth factor 1 and fibroblast growth factor 7, are also expressed at different stages of the hair cycle in adults, suggesting that they may play a role, not only in the development of the initial hair follicles but also in hair cycling. Whether the same proteins and signaling pathways are responsible for both folliculogenesis in utero and the onset of anagen after the first hair coat is not known. Like the VDR knockout mice, hairless (hr/hr) mice have normal hair morphogenesis; however, the hairless mice develop alopecia totalis by approximately 3 weeks of age (12). The onset of this alopecia coincident with the telogen phase of the first hair coat supports the hypothesis that factors that regulate the hair cycle postnatally are largely distinct from those responsible for hair follicle morphogenesis.

The association of VDR gene mutations with alopecia in both humans with hereditary 1,25-dihydroxyvitamin D- resistant rickets (HVDRR) (21) and mice (22, 3), combined with the observation that VDR-ablated mice develop alopecia regardless of their mineral ion status [2 and Fig. 5], suggests that mutation of the VDR per se is responsible for the hair loss. The absence of alopecia in profound vitamin D deficiency and in patients with 25-hydroxyvitamin D3 1α-hydroxylase mutations, further supports the hypothesis that the pathogenesis of the alopecia is a consequence of impaired receptor function rather than ligand deficiency. It is possible, however, that even in the absence of low or undetectable levels of circulating vitamin D metabolites, the skin is able to produce a vitamin D metabolite capable of mediating physiological effects. Therefore, absence, rather than deficiency, of ligand or functional receptor may be required for alopecia to be observed.

We have demonstrated that VDR ablation does not have significant effects on keratinocyte proliferation or differentiation in vitro or in vivo in neonatal mice. VDR ablation could, however, lead to alopecia by alternative means, including ligand independent effects of the VDR (23), hormone toxicity or interactions with an alternative receptor.

Because 1,25(OH)2D3 down-regulates its own biosynthesis, by repressing the 25-hydroxyvitamin D-1α-hydroxylase gene (24, 25) and increases its metabolism by up-regulating the 24-hydroxylase gene through VDR-dependent actions (26, 27), VDR-ablated cells, which possess these enzymes may have very high intracellular levels of 1,25(OH)2D3 even in the setting of normal mineral ion homeostasis. This hormone or its metabolites produced locally in the skin may have a toxic effect on hair growth by interacting with a membrane receptor or second nuclear receptor. The receptor mediating this toxic effect could be specific for vitamin D metabolites, or alternatively, bind an unrelated ligand under normal physiological conditions. Precedent for this latter suggestion has been provided by studies examining the interaction of progesterone with the oxytocin receptor (28). In these studies, the effect of progesterone on uterine sensitivity to oxytocin was shown to involve direct, nongenomic actions of progesterone on the rat oxytocin receptor. Like progesterone, vitamin D or its metabolites may mediate effects by interacting with other receptors. One would not expect these effects to be evident in the intact animal with a functional VDR, since the duration and degree of 1,25(OH)2D3 toxicity required would be anticipated to result in fatal hypercalcemia.

Elucidation of the molecular basis for the alopecia in the VDR-ablated mice will provide insight, not only into factors that regulate hair follicle homeostasis, but is also expected to clarify novel actions of this nuclear receptor in the skin.

Acknowledgments

We would like to express our appreciation to Dr. G. P. Dotto for advice on keratinocyte cultures and to Dr. J. P. Stoye for his gift of the cDNA for mhr.

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Keratinocyte

Primary type of cell found in the epidermis

Keratinocytes (stained green) in the skin of a mouse

Keratinocytes are the primary type of cell found in the epidermis, the outermost layer of the skin. In humans, they constitute 90% of epidermal skin cells. [1] Basal cells in the basal layer (stratum basale) of the skin are sometimes referred to as basal keratinocytes.[2]

Function[edit]

The primary function of keratinocytes is the formation of a barrier against environmental damage by heat, UV radiation, water loss, pathogenicbacteria, fungi, parasites, and viruses.

Pathogens invading the upper layers of the epidermis can cause keratinocytes to produce proinflammatory mediators, particularly chemokines such as CXCL10 and CCL2 (MCP-1) which attract monocytes, natural killer cells, T-lymphocytes, and dendritic cells to the site of pathogen invasion.[3]

Structure[edit]

A number of structural proteins (filaggrin, keratin), enzymes (proteases), lipids, and antimicrobial peptides (defensins) contribute to maintain the important barrier function of the skin. Keratinization is part of the physical barrier formation (cornification), in which the keratinocytes produce more and more keratin and undergo terminal differentiation. The fully cornified keratinocytes that form the outermost layer are constantly shed off and replaced by new cells.[4]

Cell differentiation[edit]

Epidermal stem cells reside in the lower part of the epidermis (stratum basale) and are attached to the basement membrane through hemidesmosomes. Epidermal stem cells divide in a random manner yielding either more stem cells or transit amplifying cells.[5] Some of the transit amplifying cells continue to proliferate then commit to differentiate and migrate towards the surface of the epidermis. Those stem cells and their differentiated progeny are organized into columns named epidermal proliferation units.[6]

During this differentiation process, keratinocytes permanently withdraw from the cell cycle, initiate expression of epidermal differentiation markers, and move suprabasally as they become part of the stratum spinosum, stratum granulosum, and eventually corneocytes in the stratum corneum.

Corneocytes are keratinocytes that have completed their differentiation program and have lost their nucleus and cytoplasmicorganelles.[7] Corneocytes will eventually be shed off through desquamation as new ones come in.

At each stage of differentiation, keratinocytes express specific keratins, such as keratin 1, keratin 5, keratin 10, and keratin 14, but also other markers such as involucrin, loricrin, transglutaminase, filaggrin, and caspase 14.

In humans, it is estimated that keratinocytes turn over from stem cells to desquamation every 40–56 days,[8] whereas in mice the estimated turnover time is 8–10 days.[9]

Factors promoting keratinocyte differentiation are:

Since keratinocyte differentiation inhibits keratinocyte proliferation, factors that promote keratinocyte proliferation should be considered as preventing differentiation. These factors include:

Interaction with other cells[edit]

Within the epidermis keratinocytes are associated with other cell types such as melanocytes and Langerhans cells. Keratinocytes form tight junctions with the nerves of the skin and hold the Langerhans cells and intra-dermal lymphocytes in position within the epidermis. Keratinocytes also modulate the immune system: apart from the above-mentioned antimicrobial peptides and chemokines they are also potent producers of anti-inflammatory mediators such as IL-10 and TGF-β. When activated, they can stimulate cutaneousinflammation and Langerhans cell activation via TNFα and IL-1β secretion.[citation needed]

Keratinocytes contribute to protecting the body from ultraviolet radiation (UVR) by taking up melanosomes, vesicles containing the endogenous photoprotectantmelanin, from epidermal melanocytes. Each melanocyte in the epidermis has several dendrites that stretch out to connect it with many keratinocytes. The melanin is then stored within keratinocytes and melanocytes in the perinuclear area as supranuclear “caps”, where it protects the DNA from UVR-induced damage.[27]

Role in wound healing[edit]

Wounds to the skin will be repaired in part by the migration of keratinocytes to fill in the gap created by the wound. The first set of keratinocytes to participate in that repair come from the bulge region of the hair follicle and will only survive transiently. Within the healed epidermis they will be replaced by keratinocytes originating from the epidermis.[28][29]

At the opposite, epidermal keratinocytes, can contribute to de novo hair follicle formation during the healing of large wounds.[30]

Functional keratinocytes are needed for tympanic perforation healing.[31]

Sunburn cells[edit]

A sunburn cell is a keratinocyte with a pyknoticnucleus and eosinophiliccytoplasm that appears after exposure to UVC or UVB radiation or UVA in the presence of psoralens. It shows premature and abnormal keratinization, and has been described as an example of apoptosis.[32][33]

Aging[edit]

With age, tissue homeostasis declines partly because stem/progenitor cells fail to self-renew or differentiate. DNA damage caused by exposure of stem/progenitor cells to reactive oxygen species (ROS) may play a key role in epidermal stem cell aging. Mitochondrial superoxide dismutase (SOD2) ordinarily protects against ROS. Loss of SOD2 in mouse epidermal cells was observed to cause cellular senescence that irreversibly arrested proliferation in a fraction of keratinocytes.[34] In older mice, SOD2 deficiency delayed wound closure and reduced epidermal thickness.[34]

Civatte body[edit]

A Civatte body (named after the French dermatologist Achille Civatte, 1877–1956)[35] is a damaged basal keratinocyte that has undergone apoptosis, and consist largely of keratin intermediate filaments, and are almost invariably covered with immunoglobulins, mainly IgM.[36] Civatte bodies are characteristically found in skin lesions of various dermatoses, particularly lichen planus and discoid lupus erythematosus.[36] They may also be found in graft-versus-host disease, adverse drug reactions, inflammatory keratosis (such as lichenoid actinic keratosis and lichen planus-like keratosis), erythema multiforme, bullous pemphigoid, eczema, lichen planopilaris, febrile neutrophilic dermatosis, toxic epidermal necrolysis, herpes simplex and varicella zoster lesions, dermatitis herpetiformis, porphyria cutanea tarda, sarcoidosis, subcorneal pustular dermatosis, transient acantholytic dermatosis and epidermolytic hyperkeratosis.[36]

See also[edit]

References[edit]

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  2. ^James W, Berger T, Elston D (December 2005). Andrews' Diseases of the Skin: Clinical Dermatology (10th ed.). Saunders. pp. 5–6. ISBN . Archived from the original on 2010-10-11. Retrieved 2010-06-01.
  3. ^Murphy, Kenneth (Kenneth M.) (2017). Janeway's immunobiology. Weaver, Casey (Ninth ed.). New York, NY, USA. p. 112. ISBN . OCLC 933586700.
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  16. ^Tu, CL; Oda, Y; Bikle, DD (1999). "Effects of a calcium receptor activator on the cellular response to calcium in human keratinocytes". The Journal of Investigative Dermatology. 113 (3): 340–5. doi:10.1046/j.1523-1747.1999.00698.x. PMID 10469331.
  17. ^Hennings, Henry; Michael, Delores; Cheng, Christina; Steinert, Peter; Holbrook, Karen; Yuspa, Stuart H. (1980). "Calcium regulation of growth and differentiation of mouse epidermal cells in culture". Cell. 19 (1): 245–54. doi:10.1016/0092-8674(80)90406-7. PMID 6153576. S2CID 23896865.
  18. ^Su, MJ; Bikle, DD; Mancianti, ML; Pillai, S (1994). "1,25-Dihydroxyvitamin D3 potentiates the keratinocyte response to calcium". The Journal of Biological Chemistry. 269 (20): 14723–9. doi:10.1016/S0021-9258(17)36685-1. PMID 7910167.
  19. ^Fu, G. K.; Lin, D; Zhang, MY; Bikle, DD; Shackleton, CH; Miller, WL; Portale, AA (1997). "Cloning of Human 25-Hydroxyvitamin D-1 -Hydroxylase and Mutations Causing Vitamin D-Dependent Rickets Type 1". Molecular Endocrinology. 11 (13): 1961–70. CiteSeerX 10.1.1.320.3485. doi:10.1210/me.11.13.1961. PMID 9415400.
  20. ^Kawakubo, Tomoyo; Yasukochi, Atsushi; Okamoto, Kuniaki; Okamoto, Yoshiko; Nakamura, Seiji; Yamamoto, Kenji (2011). "The role of cathepsin E in terminal differentiation of keratinocytes". Biological Chemistry. 392 (6): 571–85. doi:10.1515/BC.2011.060. hdl:2324/25561. PMID 21521076. S2CID 21148292.
  21. ^Jackson, B.; Brown, S. J.; Avilion, A. A.; O'Shaughnessy, R. F. L.; Sully, K.; Akinduro, O.; Murphy, M.; Cleary, M. L.; Byrne, C. (2011). "TALE homeodomain proteins regulate site-specific terminal differentiation, LCE genes and epidermal barrier". Journal of Cell Science. 124 (10): 1681–1690. doi:10.1242/jcs.077552. PMC 3183491. PMID 21511732.
  22. ^ abRheinwald, JG; Green, H (1975). "Serial cultivation of strains of human epidermal keratinocytes: The formation of keratinizing colonies from single cells". Cell. 6 (3): 331–43. doi:10.1016/S0092-8674(75)80001-8. PMID 1052771. S2CID 53294766.
  23. ^Truong, AB; Kretz, M; Ridky, TW; Kimmel, R; Khavari, PA (2006). "P63 regulates proliferation and differentiation of developmentally mature keratinocytes". Genes & Development. 20 (22): 3185–97. doi:10.1101/gad.1463206. PMC 1635152. PMID 17114587.
  24. ^Fuchs, E; Green, H (1981). "Regulation of terminal differentiation of cultured human keratinocytes by vitamin A". Cell. 25 (3): 617–25. doi:10.1016/0092-8674(81)90169-0. PMID 6169442. S2CID 23796587.
  25. ^Rheinwald, JG; Green, H (1977). "Epidermal growth factor and the multiplication of cultured human epidermal keratinocytes". Nature. 265 (5593): 421–4. Bibcode:1977Natur.265..421R. doi:10.1038/265421a0. PMID 299924. S2CID 27427541.
  26. ^Barrandon, Y; Green, H (1987). "Cell migration is essential for sustained growth of keratinocyte colonies: The roles of transforming growth factor-alpha and epidermal growth factor". Cell. 50 (7): 1131–7. doi:10.1016/0092-8674(87)90179-6. PMID 3497724. S2CID 21054962.
  27. ^Brenner M; Hearing VJ. (May–June 2008). "The Protective Role of Melanin Against UV Damage in Human Skin". Photochemistry and Photobiology. 84 (3): 539–549. doi:10.1111/j.1751-1097.2007.00226.x. PMC 2671032. PMID 18435612.
  28. ^Ito, M; Liu, Y; Yang, Z; Nguyen, J; Liang, F; Morris, RJ; Cotsarelis, G (2005). "Stem cells in the hair follicle bulge contribute to wound repair but not to homeostasis of the epidermis". Nature Medicine. 11 (12): 1351–4. doi:10.1038/nm1328. PMID 16288281. S2CID 52869761.
  29. ^Claudinot, S; Nicolas, M; Oshima, H; Rochat, A; Barrandon, Y (2005). "Long-term renewal of hair follicles from clonogenic multipotent stem cells". Proceedings of the National Academy of Sciences of the United States of America. 102 (41): 14677–82. Bibcode:2005PNAS..10214677C. doi:10.1073/pnas.0507250102. PMC 1253596. PMID 16203973.
  30. ^Ito, M; Yang, Z; Andl, T; Cui, C; Kim, N; Millar, SE; Cotsarelis, G (2007). "Wnt-dependent de novo hair follicle regeneration in adult mouse skin after wounding". Nature. 447 (7142): 316–20. Bibcode:2007Natur.447..316I. doi:10.1038/nature05766. PMID 17507982. S2CID 887738.
  31. ^Y Shen, Y Guo, C Du, M Wilczynska, S Hellström, T Ny, Mice Deficient in Urokinase-Type Plasminogen Activator Have Delayed Healing of Tympanic Membrane Perforations, PLOS ONE, 2012
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  33. ^Sheehan JM, Young AR (June 2002). "The sunburn cell revisited: an update on mechanistic aspects". Photochemical and Photobiological Sciences. 1 (6): 365–377. doi:10.1039/b108291d. PMID 12856704.
  34. ^ abVelarde MC, Demaria M, Melov S, Campisi J (August 2015). "Pleiotropic age-dependent effects of mitochondrial dysfunction on epidermal stem cells". Proc. Natl. Acad. Sci. U.S.A. 112 (33): 10407–12. Bibcode:2015PNAS..11210407V. doi:10.1073/pnas.1505675112. PMC 4547253. PMID 26240345.
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  36. ^ abcSeema, Chhabra; Pranay, Tanwar; Kumar, AroraSandeep (2013). "Civatte bodies: A diagnostic clue". Indian Journal of Dermatology. 58 (4): 327. doi:10.4103/0019-5154.113974. ISSN 0019-5154. PMC 3726905. PMID 23919028.

External links[edit]

Sours: https://en.wikipedia.org/wiki/Keratinocyte
EXTRAVASATION

Open Access

Peer-reviewed

  • Claudia Buerger ,
  • Nitesh Shirsath,
  • Victoria Lang,
  • Alina Berard,
  • Sandra Diehl,
  • Roland Kaufmann,
  • Wolf-Henning Boehncke,
  • Peter Wolf
  • Claudia Buerger, 
  • Nitesh Shirsath, 
  • Victoria Lang, 
  • Alina Berard, 
  • Sandra Diehl, 
  • Roland Kaufmann, 
  • Wolf-Henning Boehncke, 
  • Peter Wolf
PLOS

x

Abstract

Psoriasis is a frequent and often severe inflammatory skin disease, characterized by altered epidermal homeostasis. Since we found previously that Akt/mTOR signaling is hyperactivated in psoriatic skin, we aimed at elucidating the role of aberrant mTORC1 signaling in this disease. We found that under healthy conditions mTOR signaling was shut off when keratinocytes switch from proliferation to terminal differentiation. Inflammatory cytokines (IL-1β, IL-17A, TNF-α) induced aberrant mTOR activity which led to enhanced proliferation and reduced expression of differentiation markers. Conversely, regular differentiation could be restored if mTORC1 signaling was blocked. In mice, activation of mTOR through the agonist MHY1485 also led to aberrant epidermal organization and involucrin distribution. In summary, these results not only identify mTORC1 as an important signal integrator pivotal for the cells fate to either proliferate or differentiate, but emphasize the role of inflammation-dependent mTOR activation as a psoriatic pathomechanism.

Citation: Buerger C, Shirsath N, Lang V, Berard A, Diehl S, Kaufmann R, et al. (2017) Inflammation dependent mTORC1 signaling interferes with the switch from keratinocyte proliferation to differentiation. PLoS ONE 12(7): e0180853. https://doi.org/10.1371/journal.pone.0180853

Editor: Miroslav Blumenberg, NYU Langone Medical Center, UNITED STATES

Received: August 5, 2016; Accepted: June 6, 2017; Published: July 10, 2017

Copyright: © 2017 Buerger et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: All relevant data are within the paper and its Supporting Information files.

Funding: This work was funded by a grant to C.B. from the German Research Foundation (DFG) BU 1840/5-1 (http://dfg.de/), Germany and FWF Austrian Science Fund no. W1241 (https://www.fwf.ac.at/) to P.W. N.S. was supported by the PhD program Molecular Fundamentals of Inflammation (MOLIN) from the Medical University of Graz, Austria. W.-H.B. is supported by the Swiss National Fonds ME 9965 (http://www.snf.ch/). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

To maintain homeostasis of the healthy epidermis keratinocyte stem cells divide asymmetrically, leave the basal layer and successively develop into the spinous, granular and corneal layers, characterized by ordered expression of keratins and other marker such as involucrin, loricrin, filaggrin or transglutaminase [1]. Upon maturation, keratinocytes undergo a form of programmed cell death and are shed as corneocytes [2]. The balance between keratinocyte proliferation and differentiation is tightly regulated, but is deregulated in certain skin diseases such as psoriasis. Psoriasis is a chronic inflammatory skin disease presenting with red scaly plaques, mostly on the head, trunk and extensor sites of arms and legs [3]. These lesions are characterized by thickened, irregular stratum corneum with parakeratosis, epidermal thickening with acanthosis and absence of the granular layer. This is caused by hyperproliferating keratinocytes that are unable to properly initiate the epidermal differentiation program [4].

The molecular mediators and intracellular signaling pathways of the inflammatory psoriatic process involving Th17/Th22 cells and their effector cytokines acting on keratinocytes are well understood [4]. However, despite increasing identification of deregulated signal mediators such as STAT1 and 3, kinases of the MAPK family, PKC isoforms as well NF-kB [5–9], a comprehensive concept of the signaling pathways governing epidermal homeostasis and its alterations in diseases such as psoriasis has yet to be established.

Previously we found that inflammation dependent dysregulation of the PI3-K/Akt cascade interferes with the equilibrium between keratinocyte proliferation and differentiation and potentially contributes to the pathogenesis of psoriasis [10]. An important effector of PI3-K/Akt via TSC1/2 and the small GTPase Rheb [11] is the mTOR signaling pathway. The mTOR kinase, existing in two different multi-protein complexes (mTORC1 and 2), plays a central role in regulating cell growth and proliferation and is frequently dysregulated in different tumors [12], notably as well in epidermal tumors [13].

Active mTORC1- consisting besides the mTOR kinase itself, of Raptor and PRAS40 as well as other regulatory proteins [12]—phosphorylates proteins such as 4E-BP1 and S6 kinase 1 (S6K1), which in turn phosphorylates the ribosomal protein S6. 4E-BP1 and S6 are involved in protein biosynthesis through the regulation of translation [14]. In addition mTORC1 is able to promote through the activation of transcription factors lipid biogenesis, energy metabolism and repress autophagy [15]. The function of mTORC2 is less defined. It phosphorylates Akt at S473 and other AGC-kinases, contributes to cytoskeleton reorganization and is potentially involved in the regulation of cell tugor [16].

Our group previously reported for the first time an increase in mTOR expression and phosphorylation in psoriatic skin as well as hyperactivation of S6K1 and the ribosomal protein S6 [17]. It is currently hypothesized that the PI3-K/Akt/mTOR cascade plays a role in the pathogenesis of psoriasis by regulating the function of immune cells as well as intrinsic alterations within the epidermis (reviewed in [18]). Thus, we aimed at deciphering the contribution of mTORC1 signaling to epidermal homeostasis and its pathogenic role in the psoriatic epidermis.

We could show that in healthy keratinocytes Akt/mTOR signaling was deactivated, when differentiation was progressing. In contrast, under inflammatory conditions such as psoriasis, cytokines induced aberrant activation of the mTOR cascade. This contributed to the induction and/or maintenance of the psoriatic phenotype through the initiation of proliferation and blockade of proper differentiation, thus pointing towards mTOR as a potential target for therapeutic intervention in psoriasis.

Materials and methods

Chemicals and antibodies

All chemicals were purchased from Sigma unless stated otherwise. Rapamycin, Wortmannin, LY294002 and MHY1485 were from Calbiochem. Torin was purchased from Tocris Bioscience. Cytokines were obtained from Peprotech.

Phospho-specific (P-mTOR S2448 #5536, P-mTOR S2448 #2976, P-PRAS T246 #2997, P-S6 S235/6 #2211, P-Akt S473 #4060, P-ERk1 T202/Y204 #4370, P-p38MAPK T180/Y182 #4511, and corresponding pan antibodies (PRAS #2691, Akt #2938, S6 #2217) Rheb antibody (AP53656PU-N) was from Acris and tubulin #2128 antibody was from Cell Signaling Technology. Raptor antibody (sc27744) was from Santa Cruz, actin antibody (A1978) was from Sigma, involucrin antibody (ab20202) and β1-integrin antibody (ab30394) were from Abcam and filaggrin (PRB-417P-100) was from Convance. Ki-67 (MB67) antibody was obtained from Novus Biologicals and Keratin6 antibody (Ks6.KA12) from Thermo Scientific. Involucrin (Poly19244) for mouse IHC was from Biolegend.

Cell culture and conditions

The spontaneously immortalized human keratinocyte cell line (HaCaT) (Prof. Fusenig, Heidelberg, Germany) was cultured in DMEM (Invitrogen), 10% FCS (Biochrom), 1% penicillin/streptomycin solution (Invitrogen). NHK (normal human keratinocytes) cells were isolated from human juvenile foreskin and cultured in keratinocyte growth medium (Promo Cell) at 37°C in 5% CO2 atmosphere.

Separation of keratinocyte populations

Early passage keratinocytes were divided in KSCs, TA and PM cells on the basis of their ability to adhere to type IV collagen, as described elsewhere [19]. Briefly, cells were allowed to adhere to type IV collagen dishes for 5 min (KSC), and non-adherent cells were transferred to fresh collagen-coated dishes and allowed to attach overnight (TA). Non-adherent cells belong to the PM population. Isolation of these cells was verified by the expression of β1 integrin and keratin 6 (S1 Fig).

Differentiation of keratinocytes

To drive HaCaT cells into differentiation by post-confluent growth, increasing cell numbers (0.3 up to 6*105 cells / 12 well) were seeded. After 24h the higher cell numbers were confluent and differentiation was initiated. If not indicated otherwise, cells were harvested after another 48h. In NHK cells differentiation was induced by the addition of 2mM CaCl2 for at least 48h.

Proliferation of keratinocytes

Quantification of cell proliferation was determined by a colorimetric XTT assay (Roche) or by BrdU incorporation using a colorimetric cell proliferation ELISA (Roche). The assays were carried out according to the product instruction manual. For XTT assay, 2*104 HaCaT cells were seed in 96well plates in triplicates and treated as indicated for the individual assays. XTT reagent was usually added after 48h of treatment and absorption was measured 4h later. For BrdU assays, 1*104 cells were seeded in triplicates and after 24h treated as indicated for the individual assays. Cells were labeled over night with BrdU reagent and incorporation of BrdU was assessed the next morning.

siRNA mediated knockdown

HaCaT cells were reverse-transfected with Stealth™siRNA directed against Akt, Raptor or mTOR and BLOCK-IT™ Negative Control (Invitrogen). Briefly, 100pmol siRNA duplexes and 5μl Lipofectamine® 2000 were diluted separately in OptiMEM-I medium (Invitrogen), mixed and 6*105 HaCaT cells were added. Cells were seeded in 12 well plates with increasing cell densities according to the requirements of the following assays. NHK cells were seeded at 3*105 cells/12 well and the next day transfected with 15pmol Silencer®Select siRNA for Raptor, mTOR or control siRNA and 9 μl Lipofectamine® RNAiMAX. After another 48h cells were stimulated with 2mM CaCl2 for another 48h.

Western immunoblotting

Cells were lysed in RIPA lysis buffer (Cell Signaling Technology,), normalized, subjected to SDS–PAGE and blotted onto PVDF membranes. After blocking in 5% milk/TBS-T, membranes were probed with the indicated antibodies and visualized with HRP-conjugated secondary antibodies using ECL Substrate (Pierce). Western blots were quantified densiometrically using BioRad ImageLab software by dividing the signal intensity of the band of interest by the signal intensity of actin or tubulin bands. The results very normalized to the control and mean values of at least three independent experiments were calculated. Significant differences were determined by ANOVA and p-values are described in the figure legends. If no level of significance is given, differences were usually not significant.

Analysis of differentiation markers via quantitative RT-PCR

Total RNA was isolated using NucleoSpin RNA isolation kit (Machery&Nagel), transcribed using SuperScriptIII First-Strand Synthesis Mix (Life Technologies) and subjected to qRT-PCR using predesigned TaqMan® Gene Expression Assay probes (ThermoFisher) on an AbiPrism 7500 Fast Sequence Detector. mRNA expression was normalized to RPLPO and relative changes in the respective mRNA were quantified by the 2−ddCt method.

Immunhistochemistry

20 psoriasis patients between 18–75 years with a confirmed diagnosis of severe plaque-type psoriasis vulgaris for at least 6 month and no current systemic anti-inflammatory therapy were recruited from the clinical research department of the dermatology department of the Clinic of the Goethe-University, Frankfurt, Germany. Five healthy individuals were recruited among employees of our clinic. Written informed consent was given and the study was approved by the ethics committee of the Clinic of the Goethe-University (144/12); the Declaration of Helsinki protocols were followed. Punch biopsies (6mm) from lesional skin of patients or normal skin of healthy individuals were taken. These were cut into 8 μm cryosections, fixed in methanol or acetone (Raptor) and permeabilized with TBS-T. Specimens were blocked with 5% goat serum/TBS-T and incubated overnight at 4°C with primary or isotype antibodies. After washing, samples were incubated with AlexaFluor488 labeled secondary antibody and nuclei were stained with DAPI. Confocal images were generated using a ZeissLSM510 microscope.

Mice experiments

BALB/c mice (Charles-River) were housed in the animal facility of the Center for Medical Research, Medical University of Graz, Austria. All procedures were approved by the Austrian Government, Federal Ministry for Science and Research, (BMWF-66-010/0032-11/3b/2013) and conducted according to the NIH Guide for the Care and Use of Laboratory Animals. Mice (8–10 weeks) received MHY1485, dissolved in ethanol/propylene glycol in a ratio of 3:7 or vehicle alone topically once daily to the dorsal skin in a cumulative manner with increasing doses every third day (0.1 mg/ml, 0.3 mg/ml, 1 mg/ml, 3 mg/ml, 10 mg/ml). Double skin-fold thickness (DSFT) was assessed daily by measuring dorsal skin with a spring-loaded engineer’s micrometer (Mitutoyo). Mice were sacrificed 48h after the highest dose was applied. Mice were euthanized with an overdose of isoflurane and all efforts were made to minimize suffering. Blood serum was collected and stored at -80˚C for later use. Approximately 1 cm2 of central dorsal skin per mouse was excised, fixed immediately in 4% buffered formaldehyde, paraffin embedded and sectioned for H&E staining. Images were acquired by using a DP71 digital camera (Olympus, Melville, NY) attached to an Olympus BX51 microscope. Epidermal hyperplasia was monitored by counting epidermal cell layers at five randomly selected consecutive microscopic fields (at final magnification, x200). For quantification of epidermal thickness, five randomly selected measurements per H&E-stained cross-section of dorsal skin from each mouse were performed. All measurements were performed in a blinded manner. Results were first averaged per mouse and then averaged per treatment group for statistical analysis.

For immunhistochemistry staining paraffin sections were processed routinely. Primary antibody was applied overnight after pretreatment with Dako Retrieval solution of pH 6. Dako REAL Detection System (HRP/AEC, Rabbit/ Mouse) was used for detection, according to the manufacturer’s instructions. Images were acquired by using a DP71 digital camera (Olympus) and an Olympus BX51 microscope.

Bead immunoassay

Mouse serum cytokine and chemokine levels were measured with Mouse Cytokine/Chemokine bead immunoassay kit, ProcartaPlex, 26Plex from Affymetrix eBioscience according to the manufacturer’s specifications using the Bio-Plex 20 (Bio-Rad) and analyzed with five parametric curve fitting.

Results

mTOR signaling is deactivated during keratinocyte differentiation

To analyze the contribution of mTOR signaling to keratinocyte maturation, we used two different models of keratinocyte differentiation. HaCaT cells were driven into differentiation by post-confluent growth and differentiation was measured by the expression of involucrin and filaggrin. At the same time the mTOR pathway was shut off as measured by the phosphorylation of the mTOR kinase itself, PRAS40, 4E-BP1 and the ribosomal protein S6 (Fig 1A). However, other proliferative pathways such as Akt or Erk1 were also turned off. This was not a general phenomenon, as p38MAPK remained active (Fig 1A). In NHK (normal human keratinocytes) differentiation was induced by the addition of 2mM CaCl2, which led to the expression of involucrin beginning between 12 and 18h resulting into a strongly differentiated state at later time points. (Fig 1B). This was also seen on the RNA level by the induction of differentiation markers of different maturation stages (keratin1, involucrin, loricrin, transglutaminase and filaggrin) within a similar period (S1 Fig). Interestingly Ca2+ initially activated mTOR signaling as measured by phosphorylation of S6, but as cells became further differentiated, starting at 18h, activation of Akt and S6 started to decline (Fig 1B). Thus we hypothesize that deactivation of mTOR signaling is important for progression of keratinocyte differentiation.

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Fig 1. mTOR signaling is deactivated during differentiation and only partially contributes to the control of proliferation.

(a) Increasing densities of HaCaT were seeded to promote differentiation and harvested after 72h. Protein lysates were subjected to SDS-PAGE and Western Blotting with the indicated antibodies. (b) NHK were serum-starved and differentiation was induced with 2mM CaCl2 for the indicated time points. Protein lysates were subjected to SDS-PAGE and Western Blotting with the indicated antibodies was performed. Below a densiometrical quantification of involucrin and P-S6 levels of n = 4–5 similar blots is shown. Statistical significance was calculated with one-way ANOVA and Bonferroni multiple comparison (* p ≤0.05, ***p ≤0.001). (c) Keratinocytes stem cells (KSC), transient amplifying (TA) and postmitotic (PM) cells were separated according to their ability to adhere to type IV collagen. Protein lysates were subjected to SDS-PAGE and Western blotting with the indicated antibodies, showing that mTORC1 signaling is mainly present in undifferentiated cells. (d) Normal human skin was stained with P-mTOR S2448 and Ki-67 antibody. Nuclei were stained with DAPI. Single color and overlay images are presented, which show that mTOR is activated in proliferation cells of the basal layer. Bars represent 100 μm. (e) HaCaT cells were reverse-transfected with siRNA targeting Akt, Raptor or control siRNA and seeded in 96 well plates. After 48h proliferation was quantified using the XTT-based assay. Graph presents mean ± SEM (n = 4–8). Statistical significance was calculated with one-way ANOVA and Bonferroni multiple comparison (****p ≤0.0001, ns: non-significant). (f) To control for the efficiency of knockdown, HaCaT cells were transfected as in (e) and seeded in 6 well plates. Protein lysates were harvested after 48h and a Western blot was performed with the indicated antibodies. To show the consequences of Akt and Raptor knockdown, phosphorylation of the downstream targets GSK-3 and S6 was also captured. (g) HaCaT cells were seeded in 96 well-plates in triplicates and after 24h 50 μM LY294002, 100 nM Rapamycin or 250 nM Torin or solvent (DMSO) were added. After another 48h cell proliferation was measured with a BrdU assay. Graph presents mean ± SEM (n = 6). Statistical significance was calculated with one-way ANOVA and Bonferroni multiple comparison (****p ≤0.0001, ns: non-significant). (h) To show efficiency of the used inhibitors, HaCatT cells were treated as in (g). Protein lysates were prepared and a Western blot was performed with the indicated antibodies.

https://doi.org/10.1371/journal.pone.0180853.g001

mTOR signaling plays a minor role in regulating keratinocyte proliferation

To verify whether the shutdown of mTOR signaling is part of the differentiation process in the normal epidermis, we separated keratinocyte stem cells (KSC), transient amplifying (TA) and post-mitotic (PM) cells from primary human keratinocytes. While KSCs of the basal epidermal layer expressed low amounts of the differentiation marker involucrin, they showed high mTOR activity as measured by S6 phosphorylation (Fig 1C, S2 Fig). TA cells, that are about to leave the basal layer for differentiation, started to downregulate PI3-K signaling as measured by reduced Akt phosphorylation. In contrast, PM cells that are determined for terminal differentiation completely shut down their mTOR signaling and only little S6 phosphorylation could be detected in these cells (Fig 1C, S2 Fig). As these cells with high mTOR activity (KSC and TA cells) only represent a small proportion of the epidermis and we did not see active mTOR signaling in healthy skin before [17], we used a different staining protocol. We found that mTOR was active in certain cells of the basal layer and that some of these cells were also positive for Ki-67 (Fig 1D). Hence, we asked whether mTOR plays a role in regulating cell proliferation. Blocking Akt, as an upstream regulator of mTORC1 activity with either siRNA knockdown or LY294002 (Fig 1F abd 1H), impeded keratinocyte proliferation (Fig 1E and 1G). However, inhibition of mTORC1 signaling with rapamycin or Raptor knockdown (Fig 1F and 1H) had hardly any effect on keratinocyte proliferation (Fig 1E and 1G, S3 Fig). Noteworthy blocking both mTOR complexes with Torin1 and thus blocking Akt (Fig 1H) had a strong inhibitory effect on proliferation (Fig 1G). This argues that Akt regulates keratinocyte proliferation mainly via other pathways, while mTOR might have other functions in regulating the switch from proliferation to differentiation.

Inactivation of mTOR signaling is important for keratinocyte differentiation

To investigate this, mTORC1 signaling was inhibited in both cellular differentiation models. Rapamycin treatment enhanced the expression of differentiation markers: The expression of involucrin was increased on the protein level (Fig 2A and 2B), and enhanced expression of the mRNA of more differentiation markers such as involucrin, loricrin and filaggrin was detected (Fig 2C). Similar results were obtained when mTORC1 signaling was inhibited by siRNA knockdown of Raptor (Fig 2D and 2E). The expression of involucrin was increased in Raptor-deficient cells compared to control cells at all stages of differentiation, however a significant increase could be only measured for the highest cell density. Thus, we assume that mTOR signaling has to be below a critical level for differentiation to progress. In Raptor deficient cells, this can be achieved at an earlier time point than in control cells. However, blocking the mTOR kinase itself using either Torin or mTOR siRNA had the opposite effect by reducing the expression of involucrin (Fig 2A, 2B, 2D and 2E), which we assume is due to the role of mTORC2 in phosphorylating Akt, as Akt knock-down also impeded differentiation (S4 Fig). Thus, we argue that inactivation of mTORC1 signaling is a prerequisite for the progression of differentiation.

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Fig 2. Inhibition of mTORC1 signaling promotes differentiation.

(a) Increasing cell numbers of HaCaT cells were seeded and after 24h 100 nM Rapamycin, 250 nM Torin or solvent control (DMSO) were added and differentiation was allowed to proceed for 72h. Protein lysates were subjected to Western blotting and proteins were detected with the indicated antibodies. Quantification of 3–6 similar blots is depicted below. Statistical significance was calculated with two-way ANOVA and Bonferroni multiple comparison, comparing different treatments with the corresponding cell density in the control group (*p ≤0.05, ***p ≤0,001, ns: non-significant). (b) In NHK cells differentiation was induced with 2mM CaCl2 in the presence of 100nM Rapamycin, 250 nM Torin or solvent control for 48h. Protein lysates were subjected to Western blotting and proteins were detected with the indicated antibodies. Below a quantification of six similar blots is shown. Statistical significance was calculated with one-way ANOVA and Bonferroni multiple comparison (*p ≤0.05). (c) NHK cells were treated with 2 mM CaCl2 and 100nM Rapamycin or DMSO control. RNA was isolated and quantitative RT-PCR was performed to measure expression of the indicated differentiation markers. Graph presents mean ± SEM (n = 5–8). Statistical significance was calculated with two-way ANOVA and Bonferroni multiple comparison. (*p ≤0.05, ****p ≤0.0001, ns: non-significant). (d+e) HaCaT (d) or NHK (e) cells were transfected with siRNA specific for Raptor (Rap), mTOR or an siRNA control (si contr) and differentiation was induced either by post-confluent growth (d) or with 2mM CaCl2 (e) for 48h. Protein lysates were analyzed by Western blotting with the indicated antibodies. Below each blot a quantification of IVL and P-S6 band relative to actin bands of 4–6 similar blots is shown. Statistical significance was calculated with 2-way ANOVA and Bonferroni multiple comparison, comparing Raptor or mTOR knockdown with the corresponding differentiation state in the control group (*p ≤0.05, ** p ≤0,01, ****p ≤0,0001).

https://doi.org/10.1371/journal.pone.0180853.g002

mTOR signaling is highly active in lesional psoriatic skin

In psoriasis, the balance between proliferation and differentiation is strongly disturbed. To further investigate the role of mTOR signaling in psoriasis, lesional skin was stained for additional components of the mTOR complex and downstream signaling molecules. Rheb (Fig 3A) and Raptor (Fig 3C) were strongly overexpressed in lesional psoriatic skin compared to healthy skin (Fig 3B and 3D). PRAS40 showed a similar activation pattern as the mTOR kinase itself: activation over the whole epidermis of lesional skin but especially intense phosphorylation in the stratum basale (Fig 3E) and hardly any activity in healthy skin (Fig 3F). 4E-BP1 was also strongly activated in lesional skin (Fig 3G), compared to control skin (Fig 3H). In summary, hyperactivation of different components of mTOR signaling was detected in psoriatic skin with differential localization over the epidermis.

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Fig 3. mTORC1 and its downstream mediator are strongly activated in psoriatic lesions.

Punch biopsies from lesional psoriatic skin (a,c,e,g) of 20 patients and five healthy donors (b,d,f,h) were stained with antibodies for specific for Rheb, Raptor, P-PRAS40 and P-4E-BP. Nuclei were stained with DAPI. Representative overlay images from one patient and one healthy donor are shown. Bars represent 100 μm.

https://doi.org/10.1371/journal.pone.0180853.g003

Proinflammatory psoriatic cytokines activate the mTOR pathway via PI3-K signaling

When examining the mediators, that could be responsible for this strong activation of mTOR signaling, we found that IL-1β, IL-17A and TNF-α strongly activated the mTOR kinase, PRAS40 and the downstream targets of mTOR activity 4E-BP1 and the ribosomal protein S6 (Fig 4A and 4B). IL-6 and IL-22 only mediated mild activation of the mTOR pathway, while IL-17F and IL-23 did not show any effect (Fig 4A and 4B). Using PI-3K inhibitors showed that IL-1β and TNF-α dependent activation of mTOR signaling was mediated via PI3-K (Fig 4C and 4D). IL-1β, IL-17A and TNF-α were also able to activate MAPK/Erk1 signaling (Fig 4A), which only partially contributed to cytokine-dependent mTORC1 activation signaling in keratinocytes as MEK inhibition with U0126 did not fully prevent cytokine dependent activation of mTOR (Fig 4C and 4D).

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Fig 4. Psoriatic cytokines induce mTOR signaling.

Starved HaCaT (a) or NHK cells (b) were treated for 30 min with 20 ng/ml of the indicated cytokines. Serum starved HaCaT (c) or NHK cells (d) were treated for 30 min with the indicated inhibitors (1 μM Wortmannin, 50 μM LY294002, 50 μM U0126 or 100 nM Rapamycin), followed by a 30 min stimulation with IL-1 β or TNF- α (20ng/ml). Cell lysates were analyzed by Western blotting with antibodies for the indicated proteins. (e) HaCaT cells were seeded in triplicates in 96 well plates, starved overnight and treated with 50 μM LY 294002, 100 nM Rapamycin or 250 nM Torin or solvent (DMSO) as well as 20 ng/ml IL-1 β or TNF- α. After 48h cell proliferation was measured with a BrdU assay. Graph presents mean ± SEM (n = 6). Statistical significance was calculated with one-way ANOVA and Bonferroni multiple comparison (*p ≤0.05, ****p ≤0.0001). This shows that mTORC1 does not play a major role in controlling keratinocyte proliferation.

https://doi.org/10.1371/journal.pone.0180853.g004

Although mTOR signaling did not regulate keratinocyte proliferation under normal conditions, IL-1β as an mTOR activator induced cell proliferation to some degree, while TNF-α only had a small but not significant effect on proliferation (Fig 4E). However, this effect was not mediated via mTOR as rapamycin did not block proliferation. In contrast, inhibition of PI3-K with LY294002 strongly blocked proliferation. Interestingly, blocking both mTOR complexes with Torin and thereby also blocking Akt had a smaller inhibitory effect on proliferation than seen under basal conditions (Fig 1G).

In summary this data suggest that also under inflammatory conditions keratinocyte proliferation is mainly regulated via Akt and that mTOR signaling only partially contributes to this process.

Cytokine mediated induction of mTOR interferes with proper keratinocyte differentiation

As low mTORC1 signaling seems to be favorable for proper differentiation, we asked whether aberrant activation of mTORC1 by inflammatory cytokines is the reason for the differentiation defect in the psoriatic epidermis. Treating differentiating HaCaT cells with IL-1β, TNF-α and a mix of IL-1 β, TNF- α and IL-17A repressed the expression of involucrin on the protein (Fig 5A) and of keratin1, involucrin, loricrin and filaggrin on the mRNA level (Fig 5C), which was in parallel with their capacity to activate mTOR signaling (Fig 5A). In contrast, IL-17A and IL-22, which only showed mild activation of mTORC1 did not interfere with the expression of involucrin (Fig 5A). In NHK, the effect of the single cytokines on differentiation was not that prominent (Fig 5B and S5 Fig), however especially the mix was able to strongly repress the expression of all differentiation markers (Fig 5B and 5D). To verify that mTOR signaling is functionally involved in mediating this effect, we knocked out Raptor using siRNA, which specifically blocks mTORC1 signaling. Raptor knockdown not only induced upregulation of involucrin under differentiating condition as also seen in Fig 2D and 2E, but was able to partially rescue cytokine-induced repression of differentiation as measured by the expression of involucrin (Fig 5E and 5F). At least in HaCaT cells this rescue was significant when comparing Raptor knockdown cells with control cells (p ≤0.05) (Fig 5E). In contrast, we could only find a trend in NHK cells (Fig 5F), which believe is due to the greater experimental variance in these primary cells. By calculating the relative repression by the mix compared to differentiating cells in HaCaT (0.68 in si control vs 0.77 in si Raptor) we verified that there was indeed a trend towards normalization of involucrin expression in raptor knockdown cells and could exclude that this effect was only due to the general induction of involucrin by raptor knockdown. In contrast when both mTOR complexes were inhibited through knockdown of the mTOR kinase itself, the level of differentiation was lower as seen before and no rescue of cytokine induced differentiation repression could be detected (Fig 5E). Thus, we assume that psoriatic cytokines induce strong mTORC1 activity in the epidermis, which prevents proper epidermal stratification.

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Fig 5. Cytokine mediated activation of mTOR interferes with differentiation.

(a,c) HaCaT cells were seeded at proliferating (P; 0,6 *105 cells/12 well) or differentiating (Diff; 6* 105 cells/12 well) conditions and treated with 20 ng/ml of the indicated cytokines or a mix of IL-1 β, Il-17A and TNF- α. (b,d) NHK cells were treated with 2mM CaCl2 to induce differentiation and 20 ng/ml of the indicated cytokines or the mix of these cytokines. After 72h protein and RNA were isolated. (a,b) Protein samples were analyzed by Western blotting with the indicated antibodies. Below each blot quantification of n≥ 3 similar Western blots is depicted. Statistical significance was calculated with one-way ANOVA and Bonferroni multiple comparison (*p ≤0.05, **p ≤0.01). (c,d) Quantitative RT-PCR was performed to measure expression of the indicated differentiation markers. Graphs present mean ± SEM (n5). Statistical significance was calculated with two-way ANOVA and Bonferroni multiple comparison. (****p ≤0.0001). (e) HaCaT cells were reverse-transfected with siRNA specific for Raptor, mTOR or control siRNA and seeded at 0,6 *105 (P) or 6*105 (Diff) cells per 12 well. After 24h cells were treated with 20 ng of IL-1 β, IL-17A and TNF- α (Mix) and harvested after another 72h. Protein lysates were analyzed by Western blotting with the indicated antibodies. Below a quantification of 7–9 similar blots is shown. Statistical significance was calculated with two-way ANOVA and Bonferroni multiple comparison. For P-S6 statistical significant difference is shown for proliferating cells of knockdown cells compared to proliferating control cells (*p ≤0.05, **p ≤0.01, **** p ≤0.0001). (f) NHK cells were transfected with siRNA specific for Raptor or control siRNA. After 24h the cytokine mix and 2 mM CaCl2 were added. Differentiation was allowed to proceed for 48h, cells were harvested and protein lysates were analyzed by Western blotting with the indicated antibodies. Below a quantification of seven similar blots is shown. Statistical significance was calculated with two-way ANOVA and Bonferroni multiple comparison (*p ≤0.05, **p ≤0.01). Raptor knockdown rescues the cytokine induced repression of keratinocyte differentiation.

https://doi.org/10.1371/journal.pone.0180853.g005

Hyperactivation of mTORC1 signaling by MHY1485 induces a psoriasis-like skin morphology

To further verify that hyperactivation of mTORC1 is the reason for improper differentiation in psoriasis, we used the synthesized compound MHY1485 that was designed to specifically activate mTOR signaling [20]. MHY1485 induced mTORC1 signaling as shown by the phosphorylation of S6 or 4E-BP1 (Fig 6A and 6B), which was mediated by mTORC1 as pretreatment with rapamycin completely blocked the effect of MHY1485 (Fig 6A). mTORC2 signaling is not influenced as the phosphorylation of Akt at S473 remains unaltered (Fig 6A and 6B). Applying MHY1485 to the HaCaT differentiation model maintained continuous mTORC1 signaling; especially a significant increase in S6 phosphorylation could be detected during early differentiation. In untreated control cells S6 phosphorylation faded much earlier during differentiation. The continuous mTORC1 activation by the agonist impeded differentiation as measured by significantly reduced involucrin expression especially during later differentiation. However, the effect of the agonist was not strong enough to maintain S6 phosphorylation under fully differentiated conditions, resulting in residual expression of involucrin at the highest cell density (Fig 6C).

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Fig 6. Activation of mTORC1 signaling inhibits differentiation.

HaCaT cells (a) or NHK cells (b) were serum starved overnight and treated with increasing doses of MHY1485 or DMSO for 30 min If indicated cells were pre-treated with 100 nM Rapamycin for 30 min. Cells were harvested and protein lysates were analyzed by Western blotting with the indicated antibodies, showing that MHY1485 induces mTORC1 signaling. (c) Increasing numbers of HaCaT cells were seeded and driven into differentiation by post-confluent growth in the presence of the indicated concentrations of MHY1485. Protein lysates were analyzed by Western blotting with the indicated antibodies. Below a quantification of 6–8 similar Western blots is shown. Statistical significance was calculated with two-way ANOVA and Bonferroni multiple comparison (*p ≤0.05, **p ≤0.01). (d-g) MHY1485 or vehicle control was topically applied to the dorsal skin of mice for 12 consecutive days with increasing doses. (d) DSFT (Double Skin Fold Thickness) was measured before the first treatment (day1) and repeated every day. Data shown are mean values from one experiment, with n = 3 mice per treatment. Statistical significance was calculated with two-way ANOVA and Bonferroni multiple comparison (*p ≤0.05, **p ≤0.01, *** p ≤0.001, **** p ≤0.0001). (e) Representative images of H&E-stained sections from dorsal skin of a mouse of control and MHY 1485 treated groups (scale bar, 100 μM). (f) Evaluation of histological features, including number of epidermal layers and epidermal thickness in μM. Data shown are mean values of five measurements per mouse ± SEM. Statistical significance was calculated with Mann-Whitney test (**p ≤0.01). (g) Involucrin staining of vehicle control or MHY1485 treated mice. Overview images and close-ups are shown. MHY1485 induces in vivo a psoriasis like phenotype and interferes with proper differentiation (h) Hypothetical model how mTOR serves as a switch to determine the fate of keratinocytes.

https://doi.org/10.1371/journal.pone.0180853.g006

To substantiate these findings in vivo, MHY1485 was applied to the dorsal skin of mice with increasing doses. While the skin looked macroscopically nearly normal, the dorsal skin fold thickness (DSFT) was increased over the course of the experiment in MHY1485 treated animals (Fig 6D). Histological analysis revealed a psoriasis-like morphology (Fig 6E) characterized by more epidermal layers and a small but not significant acanthosis (Fig 6F). At the same time, proper differentiation was disturbed as involucrin was delocalized and also expressed in the spinous and granular layer, while in control mice it was exclusively detected in the upper granular and corneal layer (Fig 6G). However, topical application of MHY1485 did not lead to a systemic effect that resembled the situation in psoriasis as no induction of chemokines and Th1/Th17 cytokines could be detected (S6 Fig).

In summary, our data showed that in healthy skin, deactivation of mTORC1 signaling seems to be required for basal keratinocytes to progress through the epidermal maturation process. In contrast, in psoriasis overexpression of Th17 cytokines induces strong activation of mTOR signaling which prevents keratinocytes from proper differentiation in order to form the correctly stratified epidermis.

Discussion

We could show that healthy skin keratinocyte stem cells of the basal layer display high PI3-K/ mTOR activity, that correlates with Ki-67 staining and thus with their proliferative activity. This is supported by a recent study by Ding et al. that shows that epidermal loss of mTORC1 signaling results in reduced proliferation of epidermal progenitors cells in the basal layer leading to a hypoplastic epidermis during development [21]. Interestingly, we could not see a strong effect of mTOR inhibition on proliferation in vitro, which can be explained with the fact that cultivated keratinocytes represent a mixed population with a large proportion of cells, that have already left the cell cycle and are committed to differentiation.

The activity of mTOR greatly subsides when cells leave the basal layer and initiate differentiation. This seems to be necessary for proper keratinocyte maturation as blockade of mTORC1 facilitates the progression of differentiation. A possible mechanism could be the inhibition of autophagy by the mTOR complex [22]. Upon autophagic induction, phosphorylation of mTOR disappears, which is paralleled by differentiation [23]. Furthermore, differentiating keratinocytes undergo a selective form of nucleophagy, and mTORC1 signaling is critically involved in this process [24]. Interestingly, knockdown of the mTOR kinase blocked differentiation, which we assume is due to the effect of mTORC2 on Akt as knockdown of Akt also inhibited the expression of differentiation markers. This is in line with results by us [10] and others [25, 26] showing that Akt is active in the upper granular layer of health skin, where it likely contributes to nuclear degradation, which is an important step in the formation of the cornified envelope [27].

In contrast under inflammatory conditions such as in psoriasis, the vast presence of inflammatory cytokines of the TH1/17 family, constitutively activates mTORC1 signaling via PI3-K/ Akt, which can not only be seen in human psoriasis [17, 28] but also in different psoriasis mouse models and PUVA treatment significantly inactivated the mTOR pathway in vitro and in vivo [29, 30]. Interestingly, IL-22 is not only able to activate mTOR signaling but also induces the expression of mTOR mRNA [31]. Strong activation of mTORC1 in the basal compartment seems to be at least partially involved in proliferative control. This is underlined by reports that rapamycin blocks the proliferation of psoriatic keratinocytes [32].

However, the activity of mTORC1 in suprabasal layers of psoriatic skin could be indicative of a function of mTORC1 in aberrant maturation of the psoriatic epidermis. Cytokine dependent activation of mTOR signaling blocks differentiation, but can be rescued by mTORC1-specific inhibition. Keratin 6, which is upregulated within the psoriatic plaque and characteristic of hyperproliferative, cells in suprabasal layers [33] shows putative 5´TOP elements in its mRNA, which are specifically regulated by mTORC1. In addition, its expression is sensitive to rapamycin [34]. Thus, mTOR might regulate aberrant expression of keratin 6, which contributes to proliferation-associated keratinization and disturbs epidermal maturation. Another conceivable mechanism could be phosphorylation of STAT3 by mTOR [35], in the psoriatic epidermis [5]. STAT3 knock-in mice show mild epidermal hyperplasia coupled to aberrant keratinocyte differentiation with reduced expression of loricrin and filaggrin in suprabasal layers. Persistently elevated STAT3 signaling blocks keratinocyte differentiation in vitro [36] and in vivo probably by enforcing their stay in the proliferative compartment, which could be due to STAT3-dependent activation of proliferative genes such as cyclin D1 and c-Myc [37]. Moreover, a disturbed profile of autophagy expression markers could be seen in psoriatic skin, which suggests that aberrant mTOR activation in the inflamed skin inhibits proper autophagy needed for terminal differentiation [24].

We modeled this aberrant mTOR activation using the synthetic mTOR agonist MHY 1485, which induced psoriasis-like skin changes especially skin thickening and delocalization of involucrin. While involucrin is mainly localized in the granular layer in healthy human skin, it is overexpressed and mislocalized into the stratum spinosum in lesional psoriatic and therefor a sign for disturbed differentiation [38]. We assume that mTOR hyperactivation might have a dual effect on the processes leading to the psoriatic phenotype: In cells that still have the capacity to divide, mTORC1 signaling enforces proliferation contributing to the acanthosis seen in MHY1485 mice and in psoriasis. While in cells that are already determined for differentiation, the regular maturation program is blocked leading to aberrant epidermal maturation such as dislocation of involucrin.

Thus, mTOR inhibition seems a promising anti-psoriatic strategy. Remarkably, systemic rapamycin or its derivatives have been used for its immunosuppresive properties in anti-psoriatic trials alone or in combination with sub-therapeutic doses of cyclosporine and showed promising success [39–41]. However, our data support the notion that psoriasis patients could rather benefit from the topical use of mTOR inhibitors on the affected skin, which showed encouraging results in a small trial [42]. Locally applied it could not only inhibit the proliferation of psoriatic keratinocytes but also restore the epidermal differentiation defect [43].

In summary, we propose a model, where mTORC signaling serves as a switch between keratinocyte proliferation and differentiation (Fig 6H). In keratinocytes of the basal layer mTOR signaling is active and contributes to the control of proliferation while preventing differentiation. When cells leave the proliferative compartment, mTOR signaling is switched off which promotes differentiation. However, under inflammatory conditions this switch is hijacked by inflammatory cytokines, which prevents proper differentiation while promoting massive proliferation via Akt, leading to the phenotypic changes as seen in psoriasis. Thus controlling mTOR signaling might be a useful strategy to restore proper stratification of the psoriatic epidermis.

Supporting information

S1 Fig. Ca2+ dependent expression of differentiation markers over time.

NHK were serum-starved and differentiation was induced with 2mM CaCl2 for the indicated time points. RNA was isolated and quantitative RT-PCR was performed to measure expression of the indicated differentiation markers. Graph presents mean ± SEM (n = 3–7). Statistical significant difference between Ca2+ treated and control for each time point was calculated with one-way ANOVA and Bonferroni multiple comparison (***p≤0.001, ****p ≤0.0001).

https://doi.org/10.1371/journal.pone.0180853.s001

(TIF)

S2 Fig. mTOR signaling is switched off when keratinocytes mature.

Keratinocytes stem cells (KSC), transient amplifying (TA) and postmitotic (PM) cells were separated according to their ability to adhere to type IV collagen. In addition NHK were seeded in a normal cell culture dish without further separation (all). Protein lysates were subjected to SDS-PAGE and Western blotting with the indicated antibodies.

https://doi.org/10.1371/journal.pone.0180853.s002

(TIF)

S3 Fig. mTORC1 does not play a major in the proliferative control of keratinocytes.

HaCaT cells were reverse-transfected with siRNA targeting Raptor or control siRNA and seeded in 96 well plates. After 72h proliferation was quantified using a BrdU assay. Graph presents mean ± SEM (n = 6).

https://doi.org/10.1371/journal.pone.0180853.s003

(TIF)

S4 Fig. Akt Knockdown blocks differentiation.

HaCaT cells were reverse-transfected with siRNA specific for Akt or a siRNA control (si contr) and differentiation was induced by post-confluent growth for 72h. Protein lysates were analyzed by Western blotting with the indicated antibodies. Below each blot a quantification of n≥ 3 similar blots is shown. Statistical significant differences between control and knockdown cells of the same density were calculated with one-way ANOVA and Bonferroni multiple comparison (****p ≤0.0001).

https://doi.org/10.1371/journal.pone.0180853.s004

(TIF)

S5 Fig. Single cytokines are not able to interfere strongly with differentiation in NHK cells.

NHK cells were seeded and 24h later, differentiation was induced by the addition of 2 mM CaCl2 in the presence of 20 ng/ml of IL-1β, IL-17A, IL-22 or TNF- α or a mix of IL-1 β, IL-17A and TNF- α. After 72h RNA was isolated and quantitative RT-PCR was performed to measure expression of the indicated differentiation markers. Graph present mean ± SEM (n = 4–8). Statistical significant difference between Ca2+ and the cytokines was calculated with one-way ANOVA and Bonferroni multiple comparison (*p≤ 0.05, **p≤0.01, ****p ≤0.0001).

https://doi.org/10.1371/journal.pone.0180853.s005

(TIF)

S6 Fig. Topical application of MHY does not influence serum cytokine levels.

Mice were treated, as described in Fig 6D. At the end of treatment regimen, serum samples were collected and analyzed for protein expression of 26 cytokines and chemokines using multiplex bead immunoassay. IL-17, IL-23, IL-12, IL-1 β, IL-10, IL-6 and GM-CSF levels were not detectable. Data shown are from one experiment, with n = 2–3 mice per treatment group.

https://doi.org/10.1371/journal.pone.0180853.s006

(TIF)

Acknowledgments

The authors would like to thank R.Wolf for critical reading of the manuscript.

Author Contributions

  1. Conceptualization: CB PW W-HB.
  2. Formal analysis: CB NS.
  3. Funding acquisition: CB PW W-HB.
  4. Investigation: CB NS AB VL SD.
  5. Methodology: CB NS PW.
  6. Resources: CB NS PW.
  7. Validation: CB PW W-HB.
  8. Writing – original draft: CB NS PW W-HB RK.

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  • Cell differentiation 
  • Keratinocytes 
  • Cytokines 
  • Small interfering RNA 
  • Psoriasis 
  • Epidermis 
  • Cytokine therapy 
  • Gene expression 
Sours: https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0180853

Proliferation keratinocyte

Open Access

Peer-reviewed

  • Zhenxiang Wang,
  • Ying Wang,
  • Farhang Farhangfar,
  • Monica Zimmer,
  • Yongxin Zhang
  • Zhenxiang Wang, 
  • Ying Wang, 
  • Farhang Farhangfar, 
  • Monica Zimmer, 
  • Yongxin Zhang
PLOS

x

Abstract

Wound healing is primarily controlled by the proliferation and migration of keratinocytes and fibroblasts as well as the complex interactions between these two cell types. To investigate the interactions between keratinocytes and fibroblasts and the effects of direct cell-to-cell contact on the proliferation and migration of keratinocytes, keratinocytes and fibroblasts were stained with different fluorescence dyes and co-cultured with or without transwells. During the early stage (first 5 days) of the culture, the keratinocytes in contact with fibroblasts proliferated significantly faster than those not in contact with fibroblasts, but in the late stage (11th to 15th day), keratinocyte growth slowed down in all cultures unless EGF was added. In addition, keratinocyte migration was enhanced in co-cultures with fibroblasts in direct contact, but not in the transwells. Furthermore, the effects of the fibroblasts on keratinocyte migration and growth at early culture stage correlated with heparin-binding EGF-like growth factor (HB-EGF), IL-1α and TGF-β1 levels in the cultures where the cells were grown in direct contact. These effects were inhibited by anti-HB-EGF, anti-IL-1α and anti-TGF-β1 antibodies and anti-HB-EGF showed the greatest inhibition. Co-culture of keratinocytes and IL-1α and TGF-β1 siRNA-transfected fibroblasts exhibited a significant reduction in HB-EGF production and keratinocyte proliferation. These results suggest that contact with fibroblasts stimulates the migration and proliferation of keratinocytes during wound healing, and that HB-EGF plays a central role in this process and can be up-regulated by IL-1α and TGF-β1, which also regulate keratinocyte proliferation differently during the early and late stage.

Citation: Wang Z, Wang Y, Farhangfar F, Zimmer M, Zhang Y (2012) Enhanced Keratinocyte Proliferation and Migration in Co-culture with Fibroblasts. PLoS ONE 7(7): e40951. https://doi.org/10.1371/journal.pone.0040951

Editor: Giovambattista Pani, Catholic University Medical School, Italy

Received: July 26, 2011; Accepted: June 19, 2012; Published: July 20, 2012

Copyright: © 2012 Wang et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: This study was supported by Zyxell Inc. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Some experiments done for the manuscript revisions were supported by National Natural Science Foundation of China (81071563). No additional external funding was received for this study.

Competing interests: The authors have the following competing interest: Zyxell Inc. funded the study. Mr. Farhangfar was an employee of Regenetech Inc. at the time of the study and Ying Wang, Monica Zimmer and Yongxin Zhang are employees of ZYX Biotech Company. There are no patents, products in development or marketed products to declare. This does not alter the authors' adherence to all the PLoS ONE policies on sharing data and materials, as detailed online in the guide for authors.

Introduction

Wound repair and scar formation are complex and important processes in clinical care. Scar formation characterized by hypertrophy, contracture, and scar instability significantly impairs late functional and aesthetic outcomes [1]. Conventional split-thickness skin grafts, which are often widely meshed and expanded, are utilized to close large wound deficits. The consequences of scarring and contracture often require lengthy courses of revisional surgery [1]. Tissue repair is divided into an inflammatory phase, a granulation phase with synthesis of new connective tissue and epithelial wound closure, and finally a scar remodeling phase once the epidermal barrier has been re-established. In the mid- and late phases of wound healing, cellular interactions become dominated by the interplay of keratinocytes with fibroblasts, which gradually shift the microenvironment away from an inflammatory to a synthesis-driven granulation phase [2].

Untreated full-thickness wounds often heal with marked contracture and a deforming scar. It is well accepted that a wound that takes longer than 2–3 weeks to heal is at an increased risk of hypertrophic scar formation and contracture [3]. This risk can be reduced by the timely application of a split-thickness skin graft [4], and the application of a split-thickness skin graft or a dermal substitute to a full-thickness wound also inhibits the degree of contraction. This finding has been correlated with a reduction in inflammatory infiltrate [3]. The grafting of granulating wounds results in a rapid decrease in the number of myofibroblasts mediated by an apoptosis mechanism. However, graft contraction and hypertrophic scar formation remains a considerable problem even when skin grafts are applied in a timely manner. It has been estimated that more than 20% of patients with burns will develop significant hypertrophic scarring or graft contracture [3]. Therefore, we hypothesized that other factors, such as the interaction between keratinocytes and fibroblasts, may also significantly affect the wound healing process.

Wound healing is dependent on the recruitment of several cell types that appear in the wound area in a temporally- and spatially-defined manner [5]. Reepithelialization largely coincides with the recruitment of dermal fibroblasts, and it is likely that crosstalk between epidermal keratinocytes and fibroblasts is important during the rebuilding of tissue integrity [6]. The healing of extensive wounds often results in excessive scarring, disgorging, and functional impairment of the affected area [7]. This is a particular problem in the healing of large burn wounds, and it appears that early reepithelialization or coverage of the wounded area with autologous skin grafts limits excessive deposition of connective tissue. Experiments where keratinocytes were co-cultured with fibroblasts have demonstrated the establishment of paracrine loops of cytokines between the two cell types [8], [9], which is a phenomenon that may also occur in vivo to regulate cellular function [10]. It has been suggested that epidermal keratinocytes can down-regulate the production of the major matrix component of the dermis, collagen I, by fibroblasts [11], [12]. However, other studies have demonstrated that keratinocytes can be expanded by co-culturing them with human dermal fibroblasts (fibroblasts) without fetal calf serum [13]. It has also been shown that human fibroblasts can support the expansion of keratinocytes, and keratinocytes that were expanded on fibroblasts in the absence of serum tended to exhibit less differentiation than those expanded with serum [13]. To further understand these conflicting research results, the interaction of keratinocytes and fibroblasts in different culture stages and the molecular mechanisms involved were explored in this study.

Materials and Methods

Culture Media

Several different combinations of hormones and growth factors were used in our studies. For the cultivation of fibroblasts, we used RPMI1640 medium containing 10% (v/v) fetal bovine serum (FBS, Gibco-BRL, Germany), 100 U/mL penicillin, and 100 µg/mL streptomycin (pH 7.2) (Gibco-BRL, Germany). For the cultivation of keratinocytes, keratinocyte growth medium (KGM, serum-free medium, Provitro, Germany) was used with its supplements but containing no EGF except where otherwise indicated. For the stimulation mimic tests, EGF (R&D Systems, US), Recombinant human heparin-binding EGF-like growth factor (HB-EGF) (R&D Systems), IL-1α (R&D Systems), or TGF-β1 (R&D Systems) were added to KGM at a concentration of 0, 10, 20 or 40 ng/ml, respectively. Goat anti-human HB-EGF polyclonal antibody, mouse anti-IL1α monoclonal antibody, and mouse anti-TGF-β1 monoclonal antibody and their isotype controls were all purchased from Biocompare, South San Francisco, CA, USA and used at a concentration of 0.5 µl/ml for the corresponding cytokine neutralization tests. For the validation of the tests, the test values from all 4 isotype controls must be in the range of 1× standard deviation (±SD) of the means from 4 blank controls. Collagenase type II, Trypsin/EDTA, and phosphate-buffered saline (PBS) (0.15 M) were obtained from Gibco-BRL, Germany. Collagen type I from bovine skin was purchased from Sigma, USA.

Isolation and Culture of Human Dermal Fibroblasts

Human fibroblasts and keratinocytes were obtained from the dermis of juvenile human foreskin at the time of circumcision, with consent from the parents, as previously described [14], [15]. Samples were transferred to RPMI medium containing 10% (v/v) FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin and stored for up to 1 week at 4°C. Samples were washed three times in RPMI containing penicillin-streptomycin and Nystatin to remove possible pathogens and then rinsed several times with PBS to remove any blood and serum. Most of the subcutaneous fat and membranous material was removed with either a scalpel blade or a sharp pair of curved scissors. The skin was cut into strips 1 cm wide using a scalpel and the epidermis was separated from the dermis using two pairs of forceps.

For the separation of keratinocytes, the strips were placed, epidermis side up, in 0.25% trypsin and then incubated at 4°C overnight or at 37°C for 1 h. The dermis was cut into very small pieces (5 mm2) using curved scissors and the pieces were transferred into a collagenase Type II (200 IU/ml) solution in an incubator at 37°C for 2 h. The suspension was then centrifuged at 500×g for 5 min. The pellet was resuspended in culture medium. The isolated cells were counted using a hemocytometer and then seeded into culture dishes at a cell density of 2×105 cells/cm2. The cells were cultured in a CO2 incubator at 37°C and the culture medium was changed twice a week. The cells were passaged before reaching confluence by treating them with Trypsin-EDTA for 5 min, and then split into new dishes at a 1∶3 dilution [9], [14].

Isolation and Culture of Human Epidermal Keratinocytes

The epidermis obtained from the previous section was incubated in 0.25% trypsin solution at 4°C overnight or at 37°C for 1 h. The trypsinized tissue was transferred into a 15 ml centrifuge tube and vortexed gently for 2 min followed by the addition of an equal volume of FBS to inhibit the action of the trypsin. The suspension was centrifuged at 400×g for 5 min, the supernatant was discarded, and the cell pellet was resuspended in KGM culture medium. After counting the cells and determining the viability using the trypan blue exclusion method, the cells were seeded in tissue culture dishes coated with Collagen I/fibronectin/bovine serum albumin (BSA). Approximately 40% of the epidermal cells attached and began to spread out on the dish within 24–48 h. The culture medium was changed daily and the dishes were confluent by 7–10 d post-seeding [5], [8], [12].

Staining with PKH2 and PKH26

Fibroblasts and keratinocytes were stained with PKH2 and PKH26, respectively (Sigma ImmunoChemicals, US), before being used in the cultures according to the manufacturer’s instructions. Briefly, cells were suspended in 1 mL of diluent and immediately transferred into polypropylene tubes containing 1 mL of 4 µM PKH2 or PKH26 in diluent at room temperature. After 5 min incubation with frequent agitation, 2 mL of FBS was added to the suspension followed by an additional incubation for 1 min. The total volume was brought up to 8 mL with complete medium (Iscove’s modified Dulbecco’s medium [IMDM] supplemented with 10% FBS, 1× L-glutamine, and antibiotics [100 U/mL penicillin and 100 µg/mL streptomycin]), and the cells were washed three times in complete medium. After centrifugation at 400×g for 5 min, the cell pellet was resuspended in PBS and transferred to a new tube. The cells were washed two more times in PBS in the same tube, and then resuspended in complete medium and seeded for short-term culture as described below. Keratinocytes stained with PKH2 at appropriate concentrations for the experiments were seeded in 6-well plates, transwell plates, or 100×20 cell culture dishes pre-coated with collagen I/fibronectin/BSA based on different culture purposes as described above. The next day, the cells were washed with PBS and conditioned for 3 h with 10 ml DMEM/Ham’s F12 (4∶1) supplemented with 0.5% FBS. The cell density was approximately 50% at this point.

Co-culture Methods

Basic culture conditions.

For basic culture conditions, 0.15 or 0.3×106 fibroblasts were cultured per well of 6-well Falcon multi-well plates (pre-coated with Collagen I/fibronectin/BSA) with a surface area of 9.62 cm2 (BD Labware, Franklin Lakes, NJ) or in Falcon polyurethane cell culture inserts (upper chamber, 0.4 mm pore diameter) based on the requirements of the different experiments. After 48 h, 1.0 or 1.5×106 keratinocytes were seeded on top of the fibroblasts or in the upper chamber of the cell culture insert. The inserts were pre-coated with a mixture of 10 mg/ml bovine plasma fibronectin (Gibco BRL/Life Technology, Paisley, U.K.), 30 mg/ml bovine collagen I (Vitrogen, Cohesion, Palo Alto, CA), and 10 mg/ml BSA (Sigma) for 2 h at 37°C. As controls, fibroblasts or keratinocytes were cultured individually at the same time at the corresponding concentrations. The culture medium used before the establishment of the co-culture was the standard growth medium for the two cell types as described above. Upon initiation of the co-cultures, the medium was changed to KGM with supplements but without EGF, except where otherwise indicated, depending on the different experiment designs. The total volume was 4 ml (2+2 ml).

Migration assay.

All 6-well plates and coverslips were pre-coated with collagen I/fibronectin/BSA as described above. PKH2-stained fibroblasts (0.3×106 cells) were seeded on coverslips at the center of the bottom compartment of 6-well plates and cultured overnight at 37°C and 5% CO2. The next day, the medium was removed and 5 ml of PKH26-stained keratinocytes were seeded into each well at a density of 1×106 cells/well. In the K/Fp group, before keratinocytes were added, fibroblasts were treated with 1% paraformaldehyde for 5 minutes and washed 6 times with medium. After 48 hours, all cells were treated with Mytomycin C (10 µg/ml) for 3 hours, and washed 3 times to inhibit cell proliferation. Then, the coverslips were transferred into new wells, and 5 ml of new medium was added. At this time, some cultures received media containing anti-TGFβ1, anti-IL1α or anti HB-EGF. The cells were cultured for an additional 5 days, and then the coverslips were removed from the wells. The wells were washed with PBS and the cells were removed using a standard trypsin treatment procedure. The cells were resuspended in 0.5 ml PBS, counted, and then compared to cells from the wells where either fibroblasts or keratinocytes were not added or were added in the upper chambers of the Transwells. Each condition was tested in 4–6 replicate wells. The cell type and number were determined by Trypan Blue exclusion and Flow Cytometry. To validate our cover slip method, scratch assay, a classic method [16]–[20] for evaluating cell migration, was used for the correlation analysis in different concentrations of cells and HB EGF (0, 5, 10, 20 and 40/ng/ml) which resulted in different cell migration distances. Cells in scratch assay were also treated with Mitomycin C (10 ug/ml, 3 hours) and the migration distance was measured at 48 hours following culture. When 0.1, 0.3, 0.6 and 0.9×106 fibroblasts and 0.5, 1.0, 1.5 and 2.0×106 keratinocytes were respectively combined for the tests, the cell counts in cover slip assay and the cell moving distance in scratch assay at different time points correlated very well (R2>0.90, P<0.01). It can be concluded that the results of cover slip assay objectively reflect cell migration.

Proliferation test.

This assay was similar to the migration test, but no cover slip, Mitomycin C and Paraformaldehyde were used. The cells in four wells of 6-well plates for each condition were harvested as described above after 5, 10 and 15 days of culture, respectively.

Microscopy

The keratinocytes (stained with PKH26) and fibroblasts (stained with PKH2) were grown in 6-well plates and then processed as described above. Images were taken with an inverted microscope (ECLIPSE TE2000-U; Nikon) using a red and green Fluor 20x/0.45 objective (Nikon), regular light or natural light and analyzed with Image-Pro Plus software (Media Cybernetics).

Flow Cytometry Analysis

Flow cytometry was performed on trypsinized keratinocytes as described by van Erp et al. (1988), and 1% BSA was added to the wash buffer. Briefly, the keratinocytes (stained with PKH26) and fibroblasts (stained with PKH2) grown in 6-well plates were trypsinized, washed with wash buffer (PBS containing 0.5% BSA), and then fixed with 4% paraformaldehyde in PBS. The cell population was gated to exclude cell debris and analyzed with FACS CellQuest software (BD, US). Fifteen thousand events were acquired in list mode according to the following four parameters: forward scatter, side scatter, green fluorescence, and red fluorescence. PKH2 or PKH26 positive cells were calculated with the formula: total cell number (from Trypan Blue exclusion cell counting) × PKH2 or PKH26 positive percentages (from flow cytometry).

Annexin V binding assay was performed by staining cells with Annexin V-PE contained in an apoptosis detection kit, according to the manufacturer’s instructions (BD Pharmingen, San Diego, CA, USA). Binding was assessed on day 5, day 10 and day 15 as in our previous studies [21].

ELISA for EGF, HB-EGF, IL-1α and TGF-β1

Cell culture supernatants were harvested and EGF (R&D Systems), HB-EGF (Biocompare, US), IL-1α (Biocompare), and TGF-β1 (Biocompare) ELISA kits were used to detect the respective cytokines following the manufacturers’ instructions, which were similar to the procedures in our previous reports [22]–[25]. Briefly, 100 µl of the sample was added into each well of a pre-coated ELISA plate in triplicate. After the incubation and washes, the detection antibodies were added. Following additional incubation and wash steps, the substrates were added and color was developed. The absorbance in each well was read at a wavelength of 415 nm using an automatic microplate reader (Molecular Devices, Sunnyvale, CA). The data were collected using SOFTmax data reduction software (Molecular Devices). All assays were conducted with standards, and the concentrations of the above cytokines in the test samples were extrapolated from the standard curves and expressed as pg/ml.

IL-1α and TGF-β1 siRNA Transfection

Fibroblasts were transfected with small interfering RNA (siRNA) using an Amaxa Nucleofector (Gaithersburg, MD) and Basic Nucleofector Kit for Mammalian Fibroblasts (Amaxa) according to the manufacturer’s protocol. Briefly, fibroblasts were cultured to 80–90% confluence, trypsinized, and resuspended in nucleofection solution with the siRNA [either negative controls (scrambled) or IL-1α or TGF-β1 siRNA (Santa Cruz Biotechnology, Inc.)]. For the validation of the assay, the test values from all 4 negative siRNA (Santa Cruz Biotechnology, Inc.) controls must be in the range of 1× standard deviation (±SD) of the means from 4 untreated cell controls. The cells were electroporated using the Amaxa nucleofection device. For each transfection, 1.5×106 cells were electroporated with either 2 µg of the negative control, IL-1α or TGF-β1 siRNA. After the transfection, the fibroblasts were resuspended in RPMI media (GIBCO, Carlsbad, CA), counted with Trypan blue exclusion, and incubated at 37°C for 10 min. An appropriate number of cells were plated in wells of a 6-well plate and then incubated for an additional 48 h in normal culture medium (described above) prior to use.

Statistical Analysis

All results are presented as means with standard errors unless otherwise stated. ANOVA and Student’s t tests were used for the comparison of means, and the z-test was used to assess the correlation analysis. P values less than 0.05 were considered statistically significant. SAS Statview software (SAS, USA) was used for data analysis.

Results

The Growth of Keratinocytes and Fibroblasts in Co-culture

To distinguish keratinocytes from fibroblasts in the co-culture, keratinocytes were stained with PHK26 and fibroblasts were stained with PHK2. The fibroblasts (3.0×105/well), stained with PHK2 (green) were cultured in pre-coated (Collagen I/fibronectin/BSA), 6-well plates for 48 hours (Fig. 1A, C), and then keratinocytes (1×106/well), stained with PHK26 (orange) were seeded into wells with (Fig. 1C) or without (Fig. 1B) fibroblasts. Keratinocytes attached to the bottom of the well by 2 days post-seeding (Fig. 1D). After 10 days in culture, the fibroblasts were completely confluent (Fig. 1E and G), and the keratinocytes tended to cover the fibroblasts to form skin-like structures (Fig. 1G). In the first 10 days of the culture, fibroblasts (Fig. 1A, C–E) proliferated faster than the kerotinocytes (Fig. 1B–D, F), although keratinocytes were seeded at a higher density. We also observed that the keratinocytes cultured with the fibroblasts (Fig. 1G) had a higher density than those cultured alone (Fig. 1F). On day 15 after the cultures were started, the fibroblasts overgrew and formed several layers of cells when grown alone (Fig. 1H); however, in the wells where the cells were co-cultured together, keratinocytes still covered most of the fibroblasts (Fig. 1I).

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Figure 1. Keratinocytes and fibroblasts in cultures.

3×105 fibroblasts, stained with PHK2 (green), were cultured in 6-well pre-coated plate for 48 hours (1A and 1C), and then 1×106 keratinocytes, stained with PHK26 (orange), were seeded into the wells with (1C) or without (1B) fibroblasts. After 2 days, keratinocytes were also attached on the bottom of the well (1D). On day 10, fibroblasts tended to cover the entire bottom of the well (1E) but keratinocytes did not proliferate very much (1F). In the co-culture, significantly more keratinocytes (round cells) were growing on the fibroblast layer (1G). On day 15, the overgrowing fibroblasts resulted in some cells grew on others (1H), while keratinocytes could still form the skin-like layer on the top of fibroblasts (1I). 1A–1F were under fluorescence microscope (x200 or x400), 1G, 1H and 1I were under regular light microscope (x200), and natural light were used for 1I(x200).

https://doi.org/10.1371/journal.pone.0040951.g001

Interactions between Keratinocytes and Fibroblasts in the Co-culture

Keratinocytes were cultured in the following conditions: with fibroblasts in common 6-well plates (Kera/Fibr), with fibroblasts in transwell plates (Kera//Fibr), with varying concentrations of EGF (Ker + EGF), or cultured alone (Kera). On day 5 after the experiment was initiated, the cells were trypsinized and counted by Trypan blue exclusion and flow cytometry. The keratinocyte cell count in the Kera/Fibr was significantly higher (P<0.05) than Kera//Fibr and Kera, suggesting that the increase in the number of keratinocytes in the early stage of the culture is secondary to the contact between keratinocytes and fibroblasts, although such diverges cannot be seen at later time points. To mimic the effects of fibroblasts on the keratinocytes, different concentrations of EGF were added to the keratinocyte cultures. Similar to the fibroblasts, EGF promoted keratinocyte growth and exhibited a dose-dependent effect (fig. 2a).

The number of fibroblasts also increased when EGF was applied to the cultures, but they were much less sensitive than keratinocytes to the effects of EGF. However, compared to the culture of fibroblasts alone, the fibroblasts did not exhibit an increase in growth when grown with keratinocytes, suggesting the fast growing fibroblasts cannot benefit from the co-culture with keratinocytes. In fact, the fibroblasts cultured with keratinocytes in both the 6-well and transwell plates had a slightly lower cell number compared to the group without keratinocytes, though the differences were not statistically significant (P>0.05). The number of fibroblasts also increased when EGF was applied to the cultures, but they were much less sensitive than keratinocytes to the effects of EGF. These results suggested that keratinocytes have a limited effect on the growth of fibroblasts, and that EGF promotes fibroblast proliferation but is not required for fibroblast proliferation (fig. 2b).

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Figure 2. Effects of different cultures on the growth of keratinocytes (2A) and fibroblasts

(2B). 2A: Keratinocytes were cultured with fibroblasts in common 6-well plates (Kera/Fibr), with Fibr in transwell toseparate keratinocytes and fibroblasts (Kera//Fibr), with different concentrations of EGF (EGF10: EGF 10 ng/ml, EGF20: EGF 20 ng/ml and EGF40: EGF 40 ng/ml) or keratinocytes alone (Kera). Compared to Kera or Fibr//Kera, *P<0.01. Four cultures in each condition. 2B: Fibroblasts were cultured with keratinocytes in common 6-well plates (Kera/Fibr), with keratinocytes in transwell to separate keratinocytes and fibroblasts (Kera//Fibr), with different concentrations of EGF (EGF10: EGF 10 ng/ml, EGF20: EGF 20 ng/ml and EGF40: EGF 40 ng/ml) or fibroblasts alone (Fibr). Compared to the groups without star, *P<0.01. Four cultures in each condition.

https://doi.org/10.1371/journal.pone.0040951.g002

Since the fibroblast-induced increase in keratinocyte proliferation was reminiscent of the observed effects of EGF, we initially hypothesized that the EGF secreted by fibroblasts was perhaps the driving force behind the increased proliferation. However, ELISA assays with pg level sensitivity failed to detect EGF in any of the cell cultures. Instead, HB-EGF, a member of EGF family, as well as IL-1α and TGFβ1 were found in all cell cultures with keratinocytes (Fig. 3). Moreover, in cultures seeded with the same number of fibroblasts, those transfected with IL-1α and TGFβ1 siRNA produced lower levels of IL-1α and TGFβ1, respectively. Interestingly, the expression of HB-EGF in these co-cultures was also significantly reduced (both P<0.05). In contrast, TGFβ1 expression in the culture with IL-1α siRNA-transfected cells and the IL-1α expression in the culture with TGFβ1 siRNA-transfected cells did not change compared to the corresponding co-cultures without siRNA-transfected cells. Since the fibroblasts did not produce a significant level of HB-EGF in our current studies, and we did not find any report showing that fibroblasts could secrete HB-EGF, the above results suggest that the siRNA-mediated reduction in the production of IL-1 and TGF resulted in a concomitant reduction of HB-EGF in the keratinocytes co-cultured with those fibroblasts. The antibody neutralization tests for these cytokines further demonstrated the effects of IL-1α and TGFβ1 on HB-EGF levels. Moreover, the neutralization of any one of these cytokines with the respective antibody led to a decrease in the levels of the other two cytokines, with the effects of IL-1α and TGFβ1 neutralization on HB-EGF level being much greater than the effects of HB-EGF neutralization on IL-1α and TGFβ1 levels. These findings suggest that HB-EGF functions downstream of IL-1α and TGFβ1 in this system. As shown in Figure 3A and B, we found that (1) in all keratinocyte/fibroblast co-cultures without transwells (K/F), the keratinocyte concentrations correlated with HB-EGF levels (r = 0.815; P<0.01) better than IL-1α and TGFβ1 levels (P>0.05), (2) the keratinocytes in the co-culture without transwells proliferated significantly faster (P<0.01) than those in transwells (K//F), and (3) the inhibitory effects of siRNA transfection on IL-1α and TGFβ1 productions in fibroblasts led to a reduction (P<0.05)in HB-EGF production and keratinocyte proliferation, but more significant reductions were observed in the antibody neutralization tests (P<0.01), especially with the anti-HB-EGF antibody. These results suggested that the keratinocyte proliferation that was enhanced by co-culturing the cells with fibroblasts was mainly mediated by the cytokines produced by both cells, in which HB-EGF might play a central role in stimulating keratinocyte growth and could be up-regulated by IL-1α and TGFβ1 produced by fibroblasts.

In addition, we also found that the IL-1α expression level in the culture with TGFβ1 siRNA- transfected fibroblasts and the TGFβ1 expression level in the culture with IL-1α siRNA- transfected fibroblasts did not decrease, while the IL-1α expression level in the culture with the anti-TGFβ1 antibody and the TGFβ1 expression level in the culture with anti-IL-1α antibody were all significantly lower than those in the K/F group (Fig. 3A). These results suggested that a potential mutual regulation between IL-1α and TGFβ1 exists when fibroblasts and keratinocytes are cultured together.

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Figure 3. HB EGF, IL-1α and TGFβ1 levels (3a) as well as keratinocyte concentrations (3b) in different cell cultures.

3×105 fibroblasts and 106 keratinocytes, respectively stained with PHK2 and PHK26, were cultured or co-cultured for 5 days. The supernatant was collected (Fig. 3a) and HB EGF, IL-1α and TGFβ1 levels were measured with ELISA. Keratinocytes (K) were harvested and resuspended in 2 ml media, and then were counted by Trypan blue exclusion and Flow Cytometry (Fig. 3b). K/F: keratinocytes were cultured with fibroblasts in common 6-well plates; K: keratinocytes alone; F: fibroblasts alone; In the co-cultures, keratinocytes were cultured with fibroblasts in transwells to separate keratinocytes and fibroblasts (K//F), with fibroblasts transfected with IL-1α siRNA (K/Fa) or TGFβ1 siRNA (K/Fb), or with fibroblasts and anti-HB EGF (K/Fc), anti-IL-1α (K/Fd) or anti-TGFβ1 (K/Fe). *P<0.05 and **P<0.01 compared to the groups without star. Four cultures in each condition. Dotted line shows the keratinocyte seeding level.

https://doi.org/10.1371/journal.pone.0040951.g003

Since the fibroblasts did not produce significant levels of HB-EGF in the experiments described above (Fig. 3A), IL-1α and TGFβ1 or other cytokines most likely play a critical role in fibroblast-stimulated keratinocyte proliferation. However, the levels of IL-1α and TGFβ1 where the cells were co-cultured in the transwell were similar to the levels when the cells were cultured in direct contact with each other. Moreover, the stronger keratinocyte proliferation induced in co-cultures required direct contact between keratinocytes and fibroblasts. To further determine the roles of these two cytokines as well as the effect of cell-to-cell contact on keratinocyte proliferation, keratinocytes were cultured in the presence of different concentrations of HB-EGF, IL-1α and TGFβ1 and then compared to the keratinocytes/fibroblasts co-culture. Fig. 4 shows that all three cytokines stimulated kerotinocyte proliferation in a dose-dependent manner, but the level of cytokine required to reach the same level of growth stimulation found in the co-cultures was as much as 10 to 20-fold higher (15–30 ng/ml) than what was found in the co-culture media (1–2 ng/ml). Therefore, it is possible that the local concentrations of these cytokines in the cell-to-cell contact micro-environment are much higher than the overall levels in the media, which may be sufficient to affect proliferation.

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Figure 4. Effects of different cytokines on the growth of Keratinocytes.

Keratinocytes and fibroblasts were respectively stained with PHK26 and PHK2 before co-culture. Keratinocytes were cultured with fibroblasts (K/F), or with 10 ng/ml, 20 ng/ml and 40 ng/ml of HB EGF (K h10, K h20 and Kh40), IL-1α (Ki10, K i20 and K i40) or TGFβ1 (K t10, K t20 and K t40), or keratinocytes alone (K). 6-well-plates were used in the culture and cells were resuspended in 10 ml after harvested on day 5 following the culture. Four cultures in each condition. Keratinocytes were counted by Trypan Blue exclusion and Flow Cytometry.

https://doi.org/10.1371/journal.pone.0040951.g004

To understand the long-term effects of cytokines and co-culture on the proliferation of keratinocytes, some cultures were observed on the 5th, 10th, and 15th day after the cultures were initiated. As shown in Fig. 5, the keratinocytes in the co-culture with additional EGF maintained fast proliferation after day 5, while those in other cultures grew much slower. The keratinocytes in the K/F group exhibited a marked reduction in the proliferation rate following day 10. Since the keratinocytes in the culture with EGF did not exhibit a reduction in the proliferation rate, the slower keratinocyte proliferation in the K/F group during the late stage of the co-culture could not be caused by an insufficient level of basic nutrition in media.

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Figure 5. Effects of co-cultures on the growth of Keratinocytes in long term culture.

Keratinocytes were cultured with fibroblasts in common 6-well plates (K/F), with Fibroblasts in transwell to separate keratinocytes and fibroblasts (K//F), with 20ng/ml EGF (K+ EGF), with fibroblasts and anti-HB EGF (K/F + anti-HB EGF), anti-IL-1α (K/F + antiIL-1a) or anti-TGFβ1 (K/F antiTGF β1) or keratinocytes alone (K). 6-well-plates were used in the culture and cells were resuspended in 12 ml after harvested. Four cultures in each condition. Keratinocytes and fibroblasts were respectively stained with PKH26 and PKH2 before culture and counted at the end of each culture (stage) by Trypan Blue exclusion and Flow Cytometry.

https://doi.org/10.1371/journal.pone.0040951.g005

To further investigate if the reduction in the keratinocyte growth rate during the different stages of the culture was related to the levels of HB-EGF, IL-1α, or TGF-β1, the levels of these cytokines in the different cultures at different time points and the relationship between cell number and concentration of these cytokines were plotted (Fig. 6A–C). All cytokine levels (Fig. 6A) and total cell concentrations (keratinocytes + fibroblasts; Fig. 6B) increased steadily during the culture time period, but the cell concentration doubled faster than the concentration of all three cytokines after day 5, which resulted in a quick reduction in the cytokine/cell ratio during the late stage of the culture (Fig. 6C). The reduction in the concentration of cytokine per cell might explain the reduction in the keratinocyte growth rate during the late stage of the co-culture. When the data were analyzed together, we found that the fibroblasts grew faster than the keratinocytes, although the latter had a much higher cell seeding density. These data suggested that the reduction in keratinocyte proliferation in the co-cultures was at least partially due to the decreasing cytokine concentrations per cell, since the production of cytokines in the culture was slower than the proliferation of cells.

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Figure 6. The relationships between cell concentration and HB-EGF, IL-1α, and TGF-β1 concentrations.

Fibroblasts (3.0×105 cells) and keratinocytes (1.0×106 cells) were cultured in 6-well plates. The supernatant and cells that contained both keratinocytes and fibroblasts from 4 wells were harvested at each time point. The cells were then resuspended in 5 ml for counting. The cytokine concentrations were measured by ELISA. (6A) HB-EGF, IL-1α, and TGF-β1 concentrations at different time points. (6B) Total cell concentration (containing both keratinocytes and fibroblasts) at different time points. (6C) The ratio of HB-EGF, IL-1α, and TGF-β1 concentrations over the total cell concentration (×106/ml) at different time points.

https://doi.org/10.1371/journal.pone.0040951.g006

To understand how keratinocytes and fibroblasts interact during migration, several models have been developed [26], [27]. These models have employed similar methods of measuring the distance [26] of migration or migrating cell numbers [27], [28]. In our present model, coverslips with pre-seeded cells were placed in cell culture dishes or 6-well plates and then removed at the end of the culture. The cells that migrated off of the coverslips were removed by trypsinization and counted by Trypan blue exclusion and flow cytometry. This model allowed us to accurately determine the number of the cells affected by experimental factors at each time point.

As shown in Fig. 7, significantly more keratinocytes co-cultured with fibroblasts migrated off of the coverslips than the keratinocytes in the transwell controls (P<0.01), kerotinocytes alone control (P<0.01), and keratinocytes co-cultured in the presence of anti-IL-1α (P<0.05), anti-TGF-β1 (P<0.05), or anti-HB-EGF (P<0.01) antibodies. In contrast, the migration of fibroblasts was unaffected by any experimental factor. These results suggest that fibroblast-produced cytokines can enhance keratinocyte migration when the two cell types are in contact, possibly due to the effects of IL-1α and TGF-β1 produced by fibroblasts at cell contact points [29], where the fibroblasts may have accessibility for the delivery of cytokines to keratinocytes. The data also suggested that these two cytokines produced by fibroblasts may increase the migration activity of keratinocyes directly or/and indirectly by up-regulating HB-EGF.

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Figure 7. The effects of co-culture on the migration of keratinocytes and fibroblasts.

Keratinocytes (1.0×106 cells) were cultured with pre-seeded 3×105 fibroblasts in common 6-well plates (K/F), with fibroblasts in a transwell to separate keratinocytes and fibroblasts (K//F), or with fibroblasts and anti-IL-1α (K/Fa), anti-TGFβ1 (K/Fb), or anti-HB EGF (K/Fc). Keratinocytes alone (K) and fibroblasts alone (F) were cultured as controls. All cells were treated with Mitomycin C and the fibroblasts in K/Fp were also treated with 1% paraformaldehyde PBS. 2 days after keratinocytes were seeded, the coverslips were transferred to new wells for additional analysis. The 6-well-plates containing cells on coverslips were used for additional analysis. After culturing the cells for 5 days, the coverslips were removed and the cells in the wells were trypsinized and resuspended in 1 ml for counting. Six cultures were analyzed for each condition. Keratinocytes and fibroblasts were stained with PKH26 and PKH2, respectively, before culturing the cells, and then counted by Trypan Blue cell exclusion and flow cytometry after culture. *P<0.05 and **P<0.01 compared to the group without the star. Between *and **, P<0.05.

https://doi.org/10.1371/journal.pone.0040951.g007

Discussion

The dynamics of fibroblasts and keratinocytes in co-culture have been studied by several investigators to illustrate the interaction between these two cell types [30]–[36]. When keratinocytes were overlaid on fibroblasts, a skin-like structure formed in some studies [37]–[39]. A similar result was obtained in our current study. Microscopic observation of keratinocytes and fibroblasts in co-culture did not show any obvious alterations in morphology, which indicated that the co-culture conditions in our study were suitable for both cell types.

In previous studies [2], [5], [40], it has been shown that the activities of both fibroblasts and keratinocytes are down-regulated at the end of wound healing, and that fibroblast activity can be further suppressed artificially. However, such down-regulation was not evident in our study. In contrast, only keratinocytes exhibited a reduction in the growth rate during the late stage of the co-culture. In the early stage of co-culture, fibroblasts significantly enhanced the proliferation of keratinocytes. Furthermore, this enhancement was related to the contact between these two cell types, and was abolished when these two cell types were separated by transwell inserts. Since EGF has been widely studied in the clinical setting as an enhancer of wound healing and has been found to inhibit scar formation [41]–[45], we hypothesized that the up-regulatory effects of fibroblasts on keratinocyte proliferation might be mediated by EGF produced by fibroblasts. Although the effects of the fibroblasts on kerotinocytes were reproduced by adding EGF to the culture media, examination of co-culture samples showed no detectable levels of EGF by ELISA. These results led us to examine other members of the EGF family.

Many cytokines are involved in wound healing [2], [46]. Heparin-binding EGF-like growth factor (HB-EGF) is a member of the EGF family of proteins [47]. It has been shown to play an important role in wound healing, cardiac hypertrophy, heart development and heart function [48]–[50]. Shirakata et al. demonstrated that HB-EGF is the predominant growth factor involved in epithelialization in skin wound healing in vivo, and that it functions by accelerating keratinocyte migration rather than proliferation [49]. Moreover, the studies of Hashimoto et al. suggested that HB-EGF is an autocrine growth factor for human keratinocytes, and HB-EGF and TGF-alpha act not only through an auto-inductive mechanism, but also by mutual amplification [51], [52]. Marikovsky et al. found that wound fluid-derived HB-EGF is synergistic with insulin-like growth factor-I for Balb/MK keratinocyte proliferation [53], [54]. Consistent with this study, we also observed that the keratinocytes under a proper dosage of irradiation stopped proliferating but could still produce higher level of HB EGF with IL1α or/and TGF β 2 stimulations, in which HB EGF levels were higher than non-stimulation but lower than non-irradiated cells with stimulation.

IL-1α and TGFβ1 are cytokines that have been extensively studied for their roles in keratinocyte-fibroblast interactions [34], [36], [55]–[59]. However, it is not clear yet how IL-1α and TGFβ1 regulate HB-EGF in kerotinocyte proliferation and migration promoted by the co-culture with fibroblasts. In our current studies, the proliferation of keratinocytes that were in contact with fibroblasts in the early stage of the co-culture was significantly enhanced compared to cultures containing only keratinocytes. In addition, this enhancement was not seen in transwell cultures or in cultures containing an anti-HB-EGF antibody, and was remarkably reduced in the cultures with anti-IL-1α or anti-TGFβ1 antibodies. When IL-1α siRNA or TGFβ1 siRNA-transfected fibroblasts was used in the co-culture, the enhancement of keratinocyte proliferation was also significantly reduced, since the production of IL-1α and TGFβ1 by fibroblasts was inhibited. HB-EGF was also reduced in cultures with IL-1α or TGFβ1 siRNA-transfected fibroblasts and in co-cultures with anti-IL-1α or anti-TGFβ1 antibodies, indicating that the keratinocyte proliferation upregulated by IL-1α and TGFβ1 in the co-culture was partially mediated by HB-EGF. The effects of the anti-HB-EGF antibody on keratinocyte proliferation were significantly greater than those of the anti-IL-1α or anti-TGFβ1 antibodies, suggesting that HB-EGF plays a central role in the regulation of kerotinocyte growth in these co-cultures. Since siRNA can only partially or/and transiently block target cytokine production and the inhibition antibodies we used in our experiments were all over-dosage, the inhibitory effects of siRNA was always weaker than those of inhibition antibodies in our observations.

After the early stage of the co-culture, the keratinocytes in all of the culture conditions proliferated at similar rates, with the exception of the EGF control (20 µg/ml of HB EGF was also used as the positive control and it led 4-fold stronger cell proliferation). The EGF culture showed faster late stage growth than the other cultures, suggesting a reduction of cell growth factors in these cultures. However, when the concentrations of the HB-EGF, IL-1α, and TGFβ1 were examined, it was found that they continued to increase as the cells grew. Further analysis into the ratio of HB-EGF levels and cell count revealed that the increase in HB-EGF, IL-1α, and TGFβ1 levels failed to match the growth rate, resulting in lower levels of these cytokines per cell during the later stage of the culture.

Both keratinocytes and fibroblasts have very high motility in culture [60]–[69], which enables them to meet and form cell-to-cell contacts even when co-cultured at very low densities. To confirm the effects of cell contact on keratinocyte proliferation and migration that was promoted by fibroblasts and relevant cytokines, transwell plates were used to separate the two cell types in this study. The two cell types were separated into the upper and lower chambers of the transwell to eliminate direct contact with each other, and the cell type being observed was placed in the lower chamber. The data clearly showed that without direct contact, the fibroblasts and the IL-1α and TGFβ1 cytokines produced by the fibroblasts did not have a significant effect on the proliferation and migration of co-cultured keratinocytes. To further confirm the effects of IL-1α, TGFβ1 and HB-EGF on keratinocyte proliferation, these cytokines were added to keratinocyte cultures at different concentrations and compared to the kerotinocye/fibroblast co-culture. The results showed that all three cytokines were able to stimulate keratinocyte proliferation, but require a 10-fold higher concentration of the cytokines, which suggested that since the overall cytokine level of the culture was insufficient to account for the observed effects, direct contact may be required to provide a microenvironment with sufficient cytokine levels.

Cell death (necrosis and apoptosis) could be a factor that affects the evaluation of cell proliferation by cell counting. To eliminate the interference of cell death on our experimental results, we did Anexin V binding assay for some samples in addition to our routine Trypan Blue Exclusion test for all samples. Trypan Blue Exclusion was assessed on days 5, 10 and 15, when the cells were harvested. The percentages of Trypan Blue-stained cells were less than 2% on days 5 and 10, and less than 5% on day 15 in all experiments. Since those auto-detached cells might be the main factor affecting the evaluation of the cell proliferation, the numbers of floating cells at different time points were examined and correlated to those of the total cells and the live cells very well (R2 = 0.922−0.978, p<0.001−0.0005), and the percentages of floating cells in total cells were at a very narrow range (3.19–4.86%), demonstrating that the floating cells did not significantly affect the evaluation of cell proliferation. Annexin V binding assay is a very sensitive method for evaluating the very early stages of Apoptosis. Using the method described in our previous study [21], we examined the Annexin V- positive cells. Unlike the dead cell counting and floating cell counting, the kerotinocytes (22.4±3.7%, n = 6) had significantly higher Annexin V positive percentages compared to fibroblasts (17.3±2.9%, n = 6, P<0.05) on day 15 following the culture. In the co-culture, the Annexin V positive percentage (19.7±4.3%, n = 6) was slightly higher than the mean Anexin V- positive percentage (19.2±4.9%, n = 6) of fibroblasts and keratinocytes sole culture, but the difference is not significant (P>0.1). In contrast, Annexin V positive percentages in all cultures are quite similar and in a very narrow range (3.2–4.7% on day 5 and 5.1–6.9% on day 10) in established cultures. These data suggested that the increased kerotinocyte proliferation in our co-culture was not caused by the reduced cell death or/and apoptosis. Since the Annexin V-positive cells were increased quickly on day 15, it is possible that the more dead cells would be observed if the cells were cultured more than 15 days.

Although it has been well-demonstrated that both IL-1α, and TGFβ1 are HB EGF inducers, the increased HB EGF in the co-culture of this study could be also caused by the higher cell number as a result of the increased cell proliferation. Examining HB EGF production in the media from keratinocyte cultures and keratinocyte-fibroblast co-cultures revealed that the keratinocyte cell number in the single culture took 15 days to reach the level found in co-cultures after only 5 days. After the media was changed and the cells were further cultured for 24 hours, the HB EGF concentration (292±48 pg/ml, n = 4) in the single culture was only less than 60% of that (541±48 pg/ml, n = 4) in the co-culture. When IL-1α (10 µg/ml) or TGFβ1(10 µg/ml) was added into these two cultures, the 24-hour HB EGF production jumped to 416±51 pg/ml (n = 4) and 475±39 pg/ml (n = 4) in the single cultures, and 647±72 pg/ml (n = 4) and 696±39 pg/ml (n = 4) in co-cultures. These data further support that co-culture, IL-1α and TGFβ1 can affect HB EGF and the higher cell number is not required for the enhanced HB EGF production in our culture system. Even so, the effects of the stronger cell proliferation in co-culture on the enhanced HB EGF production are still not negligible.

Keratinocyte proliferation and migration are fundamentally important steps in the reepithelialization of skin wounds [70]. There are several existing models for the study of keratinocyte migration. These studies have focused on either the distance of migration or the number of cells mobilized during the migration [26], [27]. In the model we used in this experiment, the number of cells that migrated off of the coverslip was counted to provide an assessment of the overall magnitude of keratinocyte migration. This coverslip-based cell culture system is very easy to use and does not require any additional equipment. We compared this assay and another classic migration assay, scratch assay [16]–[20], and found that the results of these two methods correlated very well (R2>0.90, P<0.01). Using the coverslip assay, we demonstrated that the migration of kerotinocytes was enhanced by direct contact with fibroblasts, which correlated with the levels of HB-EGF, IL-1α, and TGFβ1 in the culture media. Moreover, this enhancement was inhibited by an anti-HB-EGF antibody and reduced by anti-IL-1α and anti-TGFβ1 antibodies when they were added to the cultures. These findings suggest that direct contact with fibroblasts is required to promote the migration of keratinocytes, and that HB-EGF, IL-1α and TGFβ1 play important roles in the interaction between keratinocytes and fibroblasts. Since mytomycin C treatment was used in the migration assay, one concern is if the mytomycin C- treated keratinocytes can still produce more HB-EGF in their responses to IL-1α and TGFβ1 stimulations. We examined the HB-EGF expression at both protein level (culture supernatant) by ELISA and mRNA level (cells) by real time reverse transcription PCR which was normalized by the housekeeping gene 36B4. On day 5 following IL-1α and TGFβ1 stimulations, mytomycin C- treated keratinocytes, similar (P>0.1) to untreated ones, produced a 3–4 folds more HB-EGF protein (p<0.05−0.01) and 5–8 folds more HB-EGF mRNA (P<0.01). This further supports that HB-EGF also plays an important role in increasing the keratinocyte migration in our co-culture system.

The literature is divided on the effects of TGFβ on the migration of kerotinocytes. One study showed TGFβ suppressed migration of keratinocytes [28], but consistent with most other studies [71]–[74], our data clearly demonstrated that both TGFβ1 and IL-1α produced by fibroblasts promoted the migration of keratinocytes. Since TGFβ has at least three subtypes (TGFβ1, TGFβ2, and TGFβ3), it is possible that the different subtypes of TGFβ have different functions. Moreover, a specific subtype of TGFβ may have been dominant in the mixed subtypes of TGFβ in the Tsuboi’s study. In addition, our data are also consistent with those from the studies by Koivisto et al., in which keratinocyte migration was shown to be induced by TGFβ1, EGF and other cytokines, and these inductions were promoted by autocrine HB-EGF [75]. In our study, the up-regulation of HB-EGF by IL-1α and TGFβ1 was further highlighted.

In summary, the up-regulation of HB-EGF by IL-1α and TGFβ1 appears to be a major driving force behind the effects of co-cultured fibroblasts on the proliferation and migration of keratinocytes. In addition, our data also support the therapeutic benefits of EGF on wound healing and scar formation.

Author Contributions

Conceived and designed the experiments: YZ ZW. Performed the experiments: ZW YW FF MZ YZ. Analyzed the data: YZ ZW. Contributed reagents/materials/analysis tools: ZW FF ZM YW YZ. Wrote the paper: YZ ZW MZ FF.

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  • Keratinocytes 
  • Fibroblasts 
  • Cytokines 
  • Small interfering RNA 
  • Wound healing 
  • Cell enumeration techniques 
  • Enzyme-linked immunoassays 
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EXTRAVASATION

KERATINOCYTE PROLIFERATION, DIFFERENTIATION, AND APOPTOSIS - DIFFERENTIAL MECHANISMS OF REGULATION BY CURCUMIN, EGCG AND APIGENIN

Sivaprakasam Balasubramanian and Richard L. Eckert

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Departments of Biochemistry and Molecular Biology, Obstetrics and Gynecology and Reproductive Biology, and Dermatology, University of Maryland School of Medicine, Baltimore, Maryland 21201

Correspondence: Richard L. Eckert, Ph.D., Professor and Chair, John F.B. Weaver Endowed Professor, Biochemistry and Molecular Biology, University of Maryland School of Medicine, 108 N. Greene Street, Baltimore, Maryland 21201, Ph: 410-706-3220, Fx: 410-706-8297, E-mail: [email protected]

The publisher's final edited version of this article is available at Toxicol Appl Pharmacol

See other articles in PMC that cite the published article.

Abstract

We have proposed that it is important to examine the impact of chemopreventive agents on the function of normal human epidermal keratinocytes, since these cells comprise the barrier that protects the body from a range of environmental insults. In this context, it is widely appreciated that cancer may be retarded by consumption or topical application of naturally-occurring food-derived chemopreventive agents. Our studies show that (−)-epigallocatechin-3-gallate (EGCG), a green tea-derived polyphenol, acts to enhance the differentiation of normal human keratinocytes as evidenced by its ability to increase involucrin (hINV), transglutaminase type 1 (TG1) and caspase-14 gene expression. EGCG also stimulates keratinocyte morphological differentiation. These actions of EGCG are mediated via activation of a nPKC, Ras, MEKK1, MEK3, p38δ-ERK1/2 signaling cascade which leads to increased activator protein 1 (AP1) and CAATT enhancer binding protein (C/EBP) transcription factor expression, increased binding of these factors to DNA, and increased gene transcription. In contrast, apigenin, a dietary flavonoid derived from plants and vegetables, and curcumin, an agent derived from turmeric, inhibit differentiation by suppressing MAPK signal transduction and reducing API transcription factor level. Curcumin also acts to enhance apoptosis, although EGCG and apigenin do not stimulate apoptosis. In addition, all of these agents inhibit keratinocyte proliferation. These findings indicate that each of these diet-derived chemopreventive agents has a profound impact on normal human keratinocyte function and that they operate via distinct and sometimes opposing mechanisms. However, all are expected to act as chemopreventive agents.

Keywords: EGCG, apigenin, curcumin, TPA, epidermis, involucrin, keratinocyte differentiation, apoptosis, chemoprevention, turmeric

Keratinocyte differentiation and human involucrin gene expression

The keratinocyte is the major cell type of the multilayered stratified squamous epithelium (the epidermis) that covers the body surface. To establish epidermal structure, keratinocytes in the basal layer undergo periodic cell division which gives rise to daughter cells that differentiate to produce the suprabasal epidermal layers (Nemes and Steinert, 1999). The epidermis provides protection against cancer-promoting agents present in the environment. UV light exposure, for example, is a major cause of human skin cancer and so identifying agents that prevent UV-dependent cancer is an important strategy (Lu et al., 2002). Consumption of dietary agents that reduce keratinocyte proliferation and enhance the conversion of pre-malignant cells to differentiated cells is expected to reduce cancer development. Thus, a major goal is to identify such agents and to understand their mechanism of action. We have focused on identifying agents that regulate the differentiation of normal human keratinocytes. This is an important effort, since the normal keratinocyte is the cell that provides the interface between the body and the environment and, as such, is exposed to a wide range of potentially carcinogenic agents.

Involucrin is an AP1 transcription factor-regulated marker of keratinocyte differentiation that is expressed in the suprabasal (late spinous/granular) layers of the human epidermis. Keratinocyte differentiating agents, including calcium and phorbol ester, activate involucrin gene expression (Eckert et al., 2004; Efimova et al., 2002; Efimova et al., 2003) via a signaling cascade that includes the novel PKC isoforms (nPKC), Ras, MEKK1, and MEK3 (Agarwal et al., 1999; Welter et al., 1995; Efimova and Eckert, 2000; Efimova et al., 1998). Activation of this cascade leads to an increase in p38δ and a decrease in ERK1/2 activity leading to an increase in the levels of AP1, Sp1 and CAATT enhancer binding protein (C/EBP) expression, increased binding of these factors to the hINV promoter, and increased hINV gene expression (Banks et al., 1998; Welter et al., 1995; Crish et al., 2002). Evidence for this cascade is provided by studies using pharmacological agents, constitutively-active and dominant-negative kinases, and kinase assays (Efimova and Eckert, 2000; Efimova et al., 1998; Efimova et al., 2002; Efimova et al., 2003). As a model to study the impact of dietary agents on keratinocyte function, we have studied the impact of a range of antioxidants on expression of involucrin (Crish et al., 1993; Crish et al., 1998; Crish et al., 2002; Eckert et al., 2004). We have also examined the impact of treatment with chemopreventive agents on keratinocyte proliferation and apoptosis. It is anticipated that an effective chemopreventive agent may enhance keratinocyte differentiation, suppress keratinocyte proliferation, enhance keratinocyte apoptosis, or produce a combination of these changes. In this brief review, we compare the regulation by (−)-epigallocatechin-3-gallate (EGCG), apigenin and curcumin on these processes.

Chemopreventive agents

Green tea polyphenols

Green tea has been reported to be effective against a number of cancers including skin, oral cavity, esophagus, stomach, lung, liver, prostate, bladder, cervix, colon, and small intestine (Stoner and Mukhtar, 1995). Green tea contains several polyphenols; however, EGCG is the most abundant polyphenol present in green tea and is believed to be responsible for most of the cancer chemopreventive properties. EGCG has been reported to induce apoptosis and promoter cell growth arrest by altering expression of cell cycle regulatory proteins, altering Bax/Bcl2 function, activating killer caspases, and suppressing nuclear factor kappa B function (Gupta et al., 2004; Khan et al., 2006). EGCG is an effective cancer preventive agent in mouse skin carcinogenesis and inhibits skin cancer cell proliferation in vitro (Katiyar et al., 2001; Katiyar et al., 1997). It prevents TPA and epidermal growth factor (EGF)-induced transformation of mouse JB6 epidermal cells via suppression of JNK phosphorylation and AP1 activation (Dong et al., 1997). It also inhibits UV-light dependent activation of c-fos gene and protein expression (Chen et al., 1999).

Apigenin

Apigenin is a dietary flavonoid found at high levels in parsley, thyme, and peppermint, and also in some herbs. Because of its potential antioxidant, anti-inflammatory, and anti-tumor properties, apigenin is a candidate cancer chemopreventive agent (Birt et al., 1997; Birt et al., 1996; Lepley et al., 1996; Ross and Kasum, 2002). Apigenin treatment inhibits cell proliferation in cancer cell types (Sarkar and Li, 2004). Apigenin treatment reduces the number and the size of skin tumors that develop in response to chemical carcinogen or UVB exposure via a mechanism that involves inhibition of ornithine decarboxylase activity (Wei et al., 1990). Apigenin also inhibits the TPA-dependent increase in c-jun and c-fos gene expression and tumor promotion in mouse skin (Huang et al., 1997c), and suppresses TPA-mediated COX-2 expression by blocking Akt signal transduction and arachidonic acid release in HaCaT cells (Van Dross et al., 2005).

Curcumin

Curcumin is an important polyphenol derived from the rhizome Curcuma longa L. Curcumin has anti-inflammatory, antioxidant, anticarcinogenic, antiviral, and antiinfectious activity (Shishodia et al., 2005). These functions are attributable to curcumin’s regulation of the function of various transcription factors including NF-kB, AP1, EGr-1 and C/EBP, and the suppression of genes encoding TNF, COX-2, chemokines and cell adhesion proteins. Curcumin suppress the proliferation of cancer cells via its effects on cell cycle, apoptosis and differentiation (Huang et al., 1988a; Huang et al., 1997b; Huang et al., 1997a; Huang et al., 1988b). Curcumin treatment has been shown to suppress skin tumor development in 7,12-dimethylbenz(a)anthracene/12-O-tetradecanoylphorbol-13-acetate treated mice (Huang et al., 1997a; Limtrakul et al., 2001; Singletary et al., 1998). Curcumin also attenuates the TPA-dependent increase in skin inflammation, hyperplasia, DNA synthesis, c-fos and c-jun protein expression, and ornithine decarboxylase (ODC) activity (Huang et al., 1997b).

Chemopreventive agents differentially regulate hINV gene expression

When we initiated these studies, the role of chemopreventive agents in regulating normal human epidermal keratinocyte function had not been extensively studied, as most investigations focus on transformed keratinocytes. However, we have argued that it is important to examine the impact of chemopreventive agent treatment on the function of normal epidermal keratinocytes, since these cells comprise the interface between the body and the environment. As such, these cells are exposed to a wide range of mutagenic environmental challenges and so it makes sense to assess whether chemopreventive agents regulate their function. For example, an agent that enhances the differentiation of normal or pre-malignant human keratinocytes, and thereby removes the cell from the proliferative pool, may provide a substantial anti-cancer advantage.

We initiated our studies to determine whether these agents regulate keratinocyte differentiation by assessing the impact of treatment on transcription of the involucrin (hINV) gene (Eckert et al., 2004). Keratinocytes were transfected with an involucrin promoter-luciferase reporter plasmid, pINV-241, in which the proximal hINV promoter is linked to luciferase (Balasubramanian et al., 2002; Balasubramanian and Eckert, 2004). Our studies showed that treatment with EGCG for 24 h activates the human involucrin promoter reporter plasmid, pINV-241. Immunoblot and RT-PCR analyses show that EGCG treatment also increases endogenous hINV protein and mRNA levels. We next investigated the effects of apigenin and curcumin on hINV expression. Hsu and coworkers recently reported that EGCG causes similar differentiation-dependent event, including increased expression of p57/KIP2, keratin 1, and filaggrin, and increased transglutaminase activity in normal human keratinocytes (Hsu et al., 2005). Our investigation of the effects of apigenin and curcumin showed that neither agent regulates hINV promoter activity. Moreover, concomitant application of either apigenin or curcumin with EGCG causes suppression of the EGCG-dependent increase in pINV-241 promoter activity. Additional studies indicate that both apigenin and curcumin inhibit EGCG-dependent activation of endogenous hINV protein expression. Taken together, these studies suggest that apigenin and curcumin can oppose the differentiation-promoting action of EGCG in cultured normal human keratinocytes.

Regulation of AP 1 transcription factor function

AP1 is known to regulate involucrin gene expression in human epidermis and in keratinocytes. Our transfection studies using progressively truncated hINV promoter segments showed that the EGCG response element is located in the proximal regulatory region (PRR) spanning nucleotides −128/−110, and mutation analysis showed that the AP1-1 site, located within this region, is required for this regulation (Balasubramanian et al., 2002; Balasubramanian and Eckert, 2004; Balasubramanian et al., 2006). Based on these studies, we concluded that EGCG activates hINV gene expression via an AP1 transcription factor-dependent pathway, and that apigenin and curcumin oppose this action (Balasubramanian et al., 2002; Balasubramanian and Eckert, 2004; Balasubramanian et al., 2006). Immunoblot studies confirm that EGCG enhances the level of the AP1 factors including Fra-1, Fra-2, c-fos, fosB, Jun-B, Jun-D and c-jun. Moreover, gel supershift analysis reveals that EGCG increases binding of Fra-1 and Jun-D to the hINV promoter AP1-1 site (Balasubramanian et al., 2002; Balasubramanian and Eckert, 2004; Balasubramanian et al., 2006).

The above studies suggest that apigenin and curcumin may inhibit differentiating agent-dependent hINV gene expression by suppressing AP1 factor function. To understand these effects, we treated keratinocytes with TPA, a keratinocyte differentiation agent that increases hINV gene expression by increasing AP1 factor level and AP1 factor binding to the hINV promoter AP1 sites (Welter et al., 1995). Both apigenin and curcumin suppress the TPA-dependent increase in the level of c-jun, junB, junD, Fra-1, Fra-2 and fos B, and this is associated with decreased AP1 factor binding to the hINV promoter AP1-1 site and reduced promoter activity (Balasubramanian et al., 2002; Balasubramanian and Eckert, 2004; Balasubramanian et al., 2006). Proteasome function has been implicated in the regulation of transcription factors in human keratinocytes (Balasubramanian and Eckert, 2004). Our recent results reveal that treatment with MG132, a proteasome inhibitor, reverses the apigenin- and curcumin-dependent reduction in AP1 transcription factor levels, suggesting that proteasome function is required for apigenin and curcumin action (Balasubramanian et al., 2006; Balasubramanian and Eckert, 2006).

Differential regulation of novel protein kinase c

As noted above, involucrin expression is controlled by a novel protein kinase c (nPKC), Ras, MEKK1, MEK3 signaling cascade which ultimately acts to increase AP1 factor level and binding to the hINV promoter AP1-1 site to activate hINV gene expression (Efimova and Eckert, 2000; Efimova et al., 1998; Efimova et al., 2002; Efimova et al., 2003; Efimova et al., 2004; Eckert et al., 2004; Welter et al., 1995). Our studies indicate that EGCG activates this pathway to increase hINV gene expression, and that this response can be inhibited using BIS-IM, an agent that inhibits all PKC isoforms (Balasubramanian et al., 2002). Additional recent results indicate that novel PKC isoforms increase hINV promoter activity, that EGCG significantly increases nPKCs-dependent activation of hINV gene expression (unpublished), and that treatment with apigenin or curcumin inhibits the nPKC-dependent hINV promoter activation. This finding is consistent with other reports showing that apigenin and curcumin inhibit PKC action (Lin et al., 1997).

We recently reported the effects of apigenin and curcumin treatment on PKCδ phosphorylation. Altered PKCδ activity in human keratinocytes is associated with changes in phosphorylation at tyrosine-311 (Eckert et al., 2004; Efimova et al., 2004; Konishi et al., 2001). Increased PKCδ-Y311 phosphorylation is positively associated with TPA treatment, and is correlated with increased expression of AP1 transcription factors and increased hINV promoter activity. Co-treatment with apigenin suppresses the TPA-dependent increase in promoter activity and phosphorylation analysis reveals that apigenin-treated cells display reduced levels of PKCδ-Y311 phosphorylation when compared to untreated cells (Balasubramanian et al., 2006). That the chemopreventive agent-dependent reduction in Y311 phosphorylation is biologically important is supported by data showing that a PKCδ tyrosine 311 mutant, PKCδ-Y311F, in which the tyrosine is converted to phenylalanine, has reduced ability to regulate AP1 factor levels and increase hINV promoter activity when compared to wild-type PKCδ (Balasubramanian et al., 2006).

Differential regulation of MAPK signaling

In human keratinocytes Ras and MEKK1 are activated in response to nPKC activation (Efimova et al., 1998). Expression of dominant-negative forms of Ras and MEKK1 suppress the EGCG-dependent increase in hINV promoter activity. This indicates that Ras and MEKK1 activity are required for the EGCG-dependent increase in hINV gene expression (Balasubramanian et al., 2002; Balasubramanian and Eckert, 2004). In contrast, apigenin and curcumin treatment inhibit the caRas- and MEKK1-dependent increase in promoter activity. Biochemical studies reveal that p38δ is activated (phosphorylated) following EGCG treatment (Balasubramanian et al., 2002), and that treatment with apigenin or curcumin suppresses this activation. Thus, EGCG and apigenin/curcumin appear to differentially regulate MAPK signaling in normal keratinocytes.

Differential regulation of cell morphology and apoptosis

We have also reported the impact of chemopreventive agent treatment on normal keratinocyte proliferation and apoptosis. Our findings indicate that EGCG, apigenin or curcumin treatment suppresses keratinocyte proliferation. Thus, in contrast to the opposing impact of EGCG versus curcumin/apigenin treatment on keratinocyte differentiation, all three compounds act to suppress proliferation. Interestingly, the reduction in cell number is associated with differential morphological changes. Normal human keratinocytes grow as loose non-structured colonies. In contrast, treatment with EGCG results in highly adherent flattened colonies, and apigenin treatment causes a circular flattened morphology. Moreover, the apigenin-associated morphology predominates in cells that are co-treated with TPA + apigenin or EGCG + apigenin (Balasubramanian et al., 2006). The apoptotic response is also varied. Treatment with EGCG or apigenin does not induce apoptosis in normal human keratinocytes (Balasubramanian et al., 2005; Balasubramanian et al., 2002; Balasubramanian and Eckert, 2004). However, curcumin causes a strong apoptotic response which is characterized by cell rounding, substrate detachment, increased number of cells with sub-G1/S DNA content, altered expression of cell cycle regulatory proteins, altered Bax/Bcl-xL expression and caspase activation in normal human keratinocytes (Balasubramanian and Eckert, 2006).

Summary

The present studies support several general conclusions regarding the impact of chemopreventive agents on normal human keratinocytes. First, these studies show that EGCG can promote keratinocyte differentiation. This is interesting, since promoting keratinocyte differentiation may be a mechanism whereby mutagenized cells can be removed from the epidermis. Second, chemopreventive agents can have opposing actions. For example, as summarized in Table 1, EGCG treatment stimulates keratinocyte differentiation but apigenin or curcumin treatment suppresses differentiation. Moreover, one chemopreventive agent may antagonize the action of another. For example, apigenin and curcumin inhibit the EGCG-dependent increase in keratinocyte differentiation (Balasubramanian and Eckert, 2004; Balasubramanian et al., 2006). This finding is perhaps not surprising, considering the structural differences among these compounds. Third, chemopreventive agents can produce different responses in normal versus immortalized/transformed keratinocytes. Thus, in normal keratinocytes EGCG increases AP1 factor levels, while in immortalized/transformed keratinocytes EGCG treatment reduces AP1 factor expression (Nomura et al., 2000; Dong et al., 1997; Barthelman et al., 1998; Chung et al., 1999). This suggests that the activity and mechanism of chemopreventive agent action may change during disease progression. Fourth, chemopreventive agents may simultaneously antagonize and synergize. Thus, while EGCG treatment increases AP1 and C/EBP factor levels in keratinocytes, this action is inhibited by curcumin and apigenin. However, in contrast, both agents act to suppress keratinocyte proliferation. This suggests that although EGCG and curcumin, for example, have opposing action on differentiation, they may still be effective when used together because of they suppress proliferation. An important lesson derived from these experiments is that not all chemopreventive agents are created equal and that it will be important to consider how simultaneous use of multiple chemopreventive agents may be used to enhance the therapeutic response.

Table 1

Differential regulation of normal human keratinocytes by chemopreventive agents

AgentEGCGApigeninCurcumin
Effective Concentration μM10 – 4010 – 2010 – 20
ProliferationSuppressesSuppressesSuppresses
DifferentiationActivatesSuppressesSuppresses
ApoptosisNoneNoneActivates
Cell MorphologyAdherent colonies & web arraysCircular flattenedSmaller, less densely packed and detached
Involucrin ExpressionActivatesInhibits differentiation agent-dependent increaseInhibits differentiation agent-dependent increase
AP1 factor level and activityIncreasesReducesReduces
C/EBP factor levelIncreasesReducesReduces
nPKC actionIncreasesSuppressesSuppresses
p38δActivatesSuppressesSuppresses
Caspase cleavage and activityNo effectNo effectActivates
Sub-G1 cellsVery fewVery fewMany

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Acknowledgments

This work utilized the facilities of the Skin Diseases Research Center of Northeast Ohio (NIH, AR39750) and was supported by a grant from the National Institutes of Health (RLE). We gratefully acknowledge our colleague Hasan Mukhtar who encouraged us to initiate studies designed to understand how diet-derived chemopreventive agents impact normal human keratinocytes.

Abbreviations

EGCG
(−)-epigallocatechin-3-gallate
TPA
12-O-tetradeconylphorbol-13-acetate
hINV
human involucrin
KSFM
keratinocyte serum-free medium
PKC
protein kinase C
nPKC
novel PKC
MAPK
mitogen-activated protein kinase
AP1
activator protein 1
C/EBP
CAATT enhancer binding protein

Footnotes

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Sours: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2698294/

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