Expression of Hypothalamic–Pituitary–Thyroid Axis Related Genes in the Human Skin

Abstract

The skin is commonly affected in thyroid diseases, but the mechanism for this association is still unclear. As the skin expresses numerous neuroendocrine elements, we tested the additional cutaneous expression of mediators operating in the hypothalamic–pituitary–thyroid axis. We found significant expression of the thyroid-stimulating hormone receptor mRNA in cultured keratinocytes, epidermal melanocytes, and melanoma cells. The presence of thyroid-stimulating hormone receptor was confirmed by northern analyses and the thyroid-stimulating hormone receptor was found to be functionally active in cyclic adenosine monophosphate signal assays. Thyroid-stimulating hormone receptor expressing cells also expressed the sodium iodide symporter and thyroglobulin genes. We also found expression of deiodinases 2 and 3 (mainly deiodinase 2) in whole skin biopsy specimens, and in the majority of epidermal and dermal cells by reverse transcription–polymerase chain reaction followed by sequencing of the amplified gene segments. There was selective expression of the gene for thyroid-stimulating hormone β; detection of the thyroid-releasing hormone gene was minimal and thyroid-releasing hormone receptor mRNA was not detected in most of the samples. Expression of functional thyroid-stimulating hormone receptor in the skin may have significant physiologic and pathologic consequences, particularly in autoimmune conditions associated with production of stimulating antibodies against the thyroid-stimulating hormone receptor. We conclude that the expanding list of neuroendocrine elements expressed in the skin supports a strong role for this system in cutaneous biology.

The skin, the largest body organ, maintains internal homeostasis by serving as a barrier between the external environment and the internal milieu. To maximize the maintenance of its physical and electrical integrity, the skin is endowed with high sensing capabilities for stressful stimuli and extremely tight spatial control over the subsequent biologic responses (Slominski and Wortsman, 2000). Those properties appear to be mediated by a local neuroendocrine response system tightly coupled to regional homeostasis (Slominski and Wortsman, 2000; Slominski et al, 2000, 2001). In this regard, fully functional elements of the hypothalamic–pituitary–adrenal axis are expressed in the skin (Slominski and Wortsman, 2000; Slominski et al, 2000, 2001).

The skin is also well recognized as a target for thyroid hormones (reviewed in Thiboutot, 1995; Slominski and Wortsman, 2000). Thus, 3,5,3′-triiodothyronine (T₃) is involved in epidermal differentiation, in enhancing local responsiveness to growth factors, in the physiology of sebaceous, eccrine and apocrine glands, and in hair growth, besides its stimulating effects on proteoglycan and glycosaminoglycan production by dermal fibroblasts (reviewed in Thiboutot, 1995; Slominski and Wortsman, 2000). These effects are probably mediated through interactions with the specific thyroid hormone receptors (TR), as TRβ and TRα mRNA are present in the skin (reviewed in Slominski and Wortsman, 2000). In thyroid dysfunction, the skin exhibits typical abnormalities in hyperthyroidism and hypothyroidism (reviewed in Thiboutot, 1995; Slominski and Wortsman, 2000). In hyperthyroidism, skin changes include erythema, palmoplantar hyperhidrosis, acropathy, and infiltrative dermopathy; also, Graves’ disease may be associated with generalized pruritus, chronic urticaria, alopecia areata, vitiligo, and diffuse skin pigmentation. In hypothyroidism, the skin is cool, dry with a pasty appearance; the epidermis is thin and hyperkeratotic; alopecia may develop, and there is diffuse myxedema. Interestingly, the generalized myxedema of hypothyroidism, but not the pretibial myxedema of Graves’ disease, is fully reversible with thyroid hormone therapy (reviewed in Thiboutot, 1995; Slominski and Wortsman, 2000).

Actual in situ production of thyroid hormones in human skin has not yet been demonstrated, although epidermal keratinocytes may be capable of deiodinating thyroxine (T₄) and T₃ (reviewed in Slominski and Wortsman, 2000). Fibroblasts of human dermal and orbital origin have been reported to express thyroid-stimulating hormone receptors (TSH-R) (Wu et al, 1996; Stadlmayr et al, 1997; Rapoport et al, 2000). Furthermore, keratinocytes have been shown to contain thyroid transcription factor-1, an important regulator of genes important in thyroid hormone formation, including the TSH-R (Suzuki et al, 1998a,b). In amphibian vertebrates (frogs), the skin produces thyroid-releasing hormone (TRH) in amounts far greater than any other neuroendocrine organ; and this cutaneous TRH reaches the pituitary to stimulate production of α-melanoctye-stimulating hormone (Vaudry et al, 1999).

We now report skin expression of molecular elements of the hypothalamic–pituitary–thyroid axis. We tested for an array of genes that included TRH,TRH-R,TSH,TSH-R, sodium iodide symporter (NIS), thyroglobulin, and deiodinases 2 and 3 (D2, D3) in skin cells, using brain, pituitary, and selected peripheral organs as appropriate controls.

MATERIALS AND METHODS

Tissues

Specimens of skin [normal and pathologic containing basal cell carcinoma (BCC)], adrenal gland, thyroid, and myometrium were collected during surgery, whereas fragments of placenta, umbilical cord, and fetal membranes were obtained after normal delivery. Pituitaries were obtained from the National Hormone and Pituitary Program, NIDDK; whole brain RNA was purchased commercially. The use of human tissues was approved by the University of Tennessee Health Science Center (UTHSC) Committee on Research Involving Human Subjects.

Cells

Normal human skin cells tested included neonatal keratinocytes, adult epidermal and follicular melanocytes and keratinocytes, and dermal and follicular papilla fibroblasts. Cells were cultured following standard protocols as previously described (Tobin and Bystryn, 1996). Established human skin-derived cell lines that were also tested included immortalized keratinocytes (HaCaT), squamous cell carcinoma (C_4–1), and human melanoma lines. The latter were established from the radial growth phase (WM 35 and SBCE2), vertical growth phase (WM 98 and WM 1341D), metastasis (WM 164) (gift of Dr M. Herlyn,Wistar Institute, Philadelphia, PA), and SKMEL188 melanoma that expresses the melanotic phenotype when cultured in Dulbecco minimal Eagle’s medium (Slominski et al, 1998). Rodent cell lines included the melanomas, hamster AbC1 and mouse S91. The culture conditions have been described in detail (Slominski et al, 1998, 1998, 1999). Rat FRTL-5 thyroid cells (ATCC CRL 8305; Interthyr Research Foundation, Baltimore, MD) were a fresh subclone (F1) that had all the functional properties previously detailed (Kohn et al, 1986; Suzuki et al, 1998a,b). Cells were diploid and between their fifth and 25th passage. They were grown in 6H medium with additions described previously (Kohn et al, 1986). To maximize TSH-R gene expression in these experiments (Saji et al, 1992), cells were maintained in 5H medium that contains no TSH for 5 d before use. Nonfunctioning FRT thyroid cells were grown in Coon’s modified F-12 medium (Sigma, ST. Louis, MO) containing 5% heat-treated, mycoplasma-free calf serum (Gibco Laboratories Life Technologies, Inc., Grand Island, NY), glutamine, and 1 mm nonessential amino acids (Suzuki et al, 1998b).

Reverse transcription–polymerase chain reaction (reverse transcription–PCR) assays

Total RNA was extracted using Trizol isolation kit (Gibco-BRL, Gaithersburg, MD). The synthesis of first-strand cDNA was performed using the Superscript preamplification system (Gibco-BRL). Five micrograms of total RNA per reaction was reverse transcribed according to the manufacturer’s protocol using oligo(dT) as the primer. Expression of genes having only one coding exon was evaluated using cDNA samples that were synthesized from DNase-treated poly(A) mRNA. Lack of DNA contamination was confirmed by negative amplification of RNA without prior reverse transcription.

All samples were standardized by the amplification of housekeeping gene GAPDH as described previously (Pisarchik and Slominski, 2001). The reaction mixture (25 μl) contained 2.5 mm MgCl₂, 2.5 mM of each deoxyribonucleoside triphosphate, 0.4 μm of each primer, 75 mm Tris–HCl (pH 8.8), 20 mm (NH₄)₂SO₄, 0.01% Tween 20, and 0.25 U of Taq polymerase (Promega, Madison, WI). The mixture was heated to 94°C for 2.5 min and then amplified for 35, 30, or 25 cycles as specified: 94°C for 30 s (denaturation), 60°C (TSHβ, TRH-R, D2, D3 genes, and first PCR round of TSH-R) or 55°C (TRH gene and second PCR round of TSH-R) for 40 s (annealing) and 72°C for 1 min (extension).

Human TSHβ (accession no. XM_002123), TSH-R (accession no. NM_000369), TRH (accession no. NM_007117), TRH-R (accession no. NM_003301), and D2 genes (accession no. AF093774) were amplified by nested PCR. Gene D3 (accession no. NM–001362) was amplified in one PCR reaction. We never amplified D3 gene for more than 25 cycles and used an appropriate amount of RNA as a control for DNA contamination for each sample. Sequences of the primers used for PCR amplifications are listed in Table I.

Table I.

List of primers used for reverse transcription–PCR amplification

Gene	Primers	Primer location	PCR product size (bp)
TSHβ	First pair of primers
	CTAACCATCAACACCACCATCTG (sense)	exon 1
	GGTTTGGTACAGTAGTTTGTCTTG (anti-sense)	exon 2
	Nested primers
	CCACCATCTGTGCTGGATATTG (sense)	exon 1
	(TATTGCACTTGCCACACTTACAG (anti-sense)	exon 2	198 bp
TSH-R	First pair of primers
	AATCCCTGTGAATGCTTTTC (sense)	exon 6
	ACTCAAGGAAAGTGGAAGTT (anti-sense)	exon 10
	Nested primers
	GTGAATGCTTTTCAGGGACTATG (sense)	exon 6
	GTCCAGGTGTTTCTTGCTATCAG (anti-sense)	exon 10	272 bp
	Primers used for direct PCR
	GGGTGCAACACGGCTGGTTT (sense)	exon 10
	CTGGGTTGTACTGCGGATTTCGG (anti-sense)	exon 10	367 bp
TRH	First pair of primers
	TTGCTGCTCGCTCTGGCTTTG (sense)	exon 2
	CTGGCGTTTTTCAGGCATCAG (anti-sense)	exon 3
	Nested primers
	CTCTTCCTCCGGGAAAACATC (sense)	exon 2
	CTCTTCTTCCCAGCTCCTTTG (anti-sense)	exon 3	354 bp
TRH-R	First pair of primers
	GCCTCTGCATTACTTACCTCCAG (sense)	exon 1
	GGTCTGACTCCTTGATGACGCTG (anti-sense)	exon 2
	Nested primers
	ACTGTATGCTCTGGTTCTTCTTG (sense)	exon 1
	GTTTCTCTGTTGGCTTCTGCTTG (anti-sense)	exon 2	567 bp
D2	First pair of primers
	TGCCTCTTCCTGGCTCTCTATG (sense)	exon 1
	CAAAGGCTACCCCGTAAGCTATG (anti-sense)	exon 2
	Nested primers
	ATGACTCGGTCATTCTGCTCAAG (sense)	exon 1
	CTGGGTACCATTGCCACTGTTG (anti-sense)	exon 2	227 bp
D3	GCCTTCATGCTCTGGCTTCTC (sense)	exon 1
	TAGTCGAGGATGTGCTGGCTC (anti-sense)	exon 1	296 bp
NIS	CTGCCCCAGACCAGTACATGCC (sense)	exons 7–8
	TGACGGTGAAGGAGCCCTGAAG (anti-sense)	exons 10–11	303 bp
Thyroglobulin	CCGCCGTCATCAGCCATGAG (sense)	exon 42
	TGAGTCCTCGCCACCCAGAGAA (anti-sense)	exon 44	395 bp

Amplification products were separated by agarose gel electrophoresis and visualized with ethidium bromide staining. The identified PCR products were excised from the agarose gel and purified by GFX PCR DNA and gel band purification kit (Amersham Pharmacia Biotech, Piscataway, NJ). PCR fragments were cloned in pGEM-T easy vector system (Promega) and purified by plasmid purification kit (Qiagen, Valencia, CA). Sequencing was performed in the Molecular Resource Center at the University of Tennessee HSC (Memphis) using Applied Biosystems 3100 Genetic Analyzer and BigDye^™ Terminator Kit (Applied Biosystems, Foster City, CA).

To detect NIS (accession no. NM_000453), thyroglobulin, and TSH-R (accession no. NM_000369) gene expression in HaCaT keratinocytes and human melanomas the RNA was purified free of genomic DNA using DNA-Free (Ambion, Austin, TX) according to recommended conditions. cDNA was synthesized from 1.0 μg of total RNA using MMLV reverse transcriptase with random-hexamer primers (Clontech, Palo Alto, CA). Polymerase chain reaction (50 μl) was performed using 3% of the total cDNA in a reaction mixture containing 10 mm Tris–HCl, pH 8.3, 50 mm KCl, 3.5 mm MgCl₂, 0.01% gelatin, 0.25 mM each deoxyribonucleoside triphosphate, 0.4 μm of each primer, 1.0 U AmpliTaq polymerase (Perkin-Elmer, Norwalk, CT), and 0.2 μg TaqStart antibody (Clontech). The primers used are in Table I. Amplifications conditions were: polymerase activation at 95°C for 5 min; 35 (thyroglobulin and TSH-R) or 40 (NIS) cycles of denaturation at 94°C for 20 s, annealing at 64°C for 1 min (thyroglobulin and TSH-R) or 70°C for 1 min (NIS), and extension at 72°C for 1 min; followed by 72°C for 7 min.

Northern blot analyses

Total RNA was prepared using a commercial kit (RNeasy Mini Kit, Qiagen). Fifteen micrograms of RNA samples were run on denatured agarose gels, blotted on Nytran membranes (Schleicher & Schuell, Keene, NH) and subjected to hybridization as described (Saji et al, 1992; Giuliani et al, 1995; Suzuki et al, 1998a,b). Full length rat or human TSH-R radiolabeled with [α-³²P]deoxycytidine triphosphate were used as probes (Tahara et al, 1991).

Cyclic adenosine monophosphate (cAMP) assays

The noted concentration of bovine TSH, in units described by the manufacturer (Sigma), or 100 μl (0.5 mg) of a standard anti-TSH-R stimulating antibodies were added with fresh medium to cell samples cultured in 24 well plates then incubated for 2 h in the presence of 0.5 mm of isobuthylmethylxanthine at 37°C in 5% CO₂. The concentrations of extracellular cAMP were measured with a nonacetylation enzyme immunoassay using the BIOTRAK cAMP enzyme immunoassay system as recommended by the manufacturer (Amersham Pharmacia Biotech). The cAMP concentration of samples was expressed as a level of stimulation relative to control incubations with no TSH or stimulating TSH antibodies.

RESULTS AND DISCUSSION

Expression of functional TSH-R in skin cells

As previous reports suggested TSH-R expression in the human skin (Wu et al, 1996; Stadlmayr et al, 1997; Rapoport et al, 2000), we tested expression of the gene by reverse transcription–PCR in a broad panel of specimens. Expression of the TSH-R gene 272 bp product of nested PCR (not shown) and the 367 bp product of direct PCR (Fig 1) was found in almost all tissues tested, including normal and pathologic skin; exceptions being the umbilical cord and myometrium (Table II). TSH-R was also detected in cultured dermal and follicular papilla fibroblasts, neonatal, follicular and immortalized HaCaT keratinocytes, squamous cell carcinoma cells, epidermal melanocytes, and in four melanoma lines. TSH-R gene expression was below detectable levels only in follicular melanocytes and in two melanoma lines (Table II). Northern blot analysis was then performed in selected human and rodent melanoma lines, and in human squamous cell carcinoma cells and immortalized epidermal keratinocytes. This revealed a TSH-R transcript of 3.3 kb, similar to a TSH-R transcript detected in control rat FRTL-5 thyroid cells (Fig 2). This transcript encoded a functional TSH-R as documented by strong responses to TSH and antibodies against TSH-R, e.g., marked stimulation of cAMP production in human melanoma, in keratinocytes, in hamster melanoma (Table III) and in mouse melanoma (not shown).

Figure 1. Expression of TSH-R, thyroglobulin, and NIS genes in cultured cells.

On the right the arrows identify reverse transcription– PCR products of TSH-R (367 bp), NIS (303 bp), and thyroglobulin (395 bp) transcripts. (1) HaCaT keratinocytes; (2–4) WM 35,WM 164, and WM 1341D, SBCE2 respectively; (5) SKMEL188 melanoma; (6) negative control no template; (7,8) normal thyroid. Left: Gibco-BRL Kb Plus ladder.

Table II.

Profile of D2, D3, TSH-R,TSH, TRH-R, and TRH genes expression in human tissues and cultured skin cells

	Genes
Specimens	TSHβ	TRH	TRH-R	D2	D3	TSH-R
Tissues
Brain (whole)	–	+	+	+	+	+
Pituitary	+	–	+	+	+	+
Skin (normal)	–0/4)	+ 1/4	–(0/4)	+ 4/4	+ 4/4	+ 3/3
Skin (involved by BCC)	–(0/3)	–(0/3)	+ 1/3	+ 3/3	+ 3/3	+ 2/2
Adrenal gland	–	+	–	+	+	+
Myometrium	+	+	–	+	+	–
Placenta	–	+	–	+	+	+
Umbilical cord	–	+	–	+	+	–
Fetal membrane	–	+	–	+	+	+
Cultured cells
Neonatal keratinocytes	–	+	–	+	+	+
Epidermal keratinocytes	+	–	–	+	+	+
Hair follicle keratinocytes	+	–	–	+	+	+
HaCaT keratinocytes	–	–	–	+	+	+
Epidermal melanocytes	–	–	–	+	–	+
Hair follicle melanocytes	+	–	–	–	–	–
Melanoma SKMEL188	+	–	+	+	–	–
Melanoma SBCE2	–	+	–	+	–	+
Melanoma WM35	–	–	–	+	–	+
Melanoma WM98	–	–	–	+	–	–
Melanoma WM164	+	–	–	+	–	+
Melanoma WM1341D	–	+	+	+	+	+
Dermal fibroblasts	–	+	–	+	+	+
Hair follicle papilla fibroblasts	–	+	–	+	+	+
Squamous cell carcinoma C1–4	+	–	–	+	–	+

In parentheses: (no. of positive/no. of biopsies tested)

Figure 2. Northern blotting detection of TSH-R gene expression in skin cells.

(A) RNA transcripts hybridizable to TSH-R cDNA; (B) ethidium bromide stain of corresponding total RNA. Following cell lines are presented: positive control represented by FRTL-5 thyroid line (1); human melanomas WM35 (2) and WM164 (7) lines; human C_1–4 squamous cell carcinoma (3) and HaCaT keratinocytes (4); hamster AbC1 melanoma (5); mouse S91 melanoma (6).

Table III.

TSH stimulation of TSH-R in hamster (AbC-1) and human (WM 164) melanoma cells and immortalized normal human keratinocytes (HaCaT) compared with thyroid cell lines (FRTL-5 and FRT) signal transduction is coupled to cAMP production

	Relative cAMP increase during 2 h incubation
Cell line	Buffer (control)	TSH (1 mU per ml)	TSH (10 mU per ml)	Anti-TSH receptor antibodies (1 mg per ml)
AbC-1	1	6.7	25.9	4.2
WM 164	1	3.6	6.1	2.8
HaCaT	1	1.5	5.2	3.2
FRTL-5	1	7.1	7.1	4.1
FRT	1	0.9	0.8	1.1

The values represent means from duplicate assays of a representative experiment. The data are presented as relative cAMP levels compared with control (addition of media alone).

The presence of TSH-R transcripts in dermal fibroblasts or in skin biopsies had previously been shown (Rapoport et al, 2000; Wu et al, 1996; Stadlmayr et al, 1997), and expression of thyroid transcription factor-1 (positive transcriptional regulator of TSH-R gene) (Shimura et al, 1994) had also been detected in keratinocytes (Suzuki et al, 1998a,b). What is novel is the detection of TSH-R in the main cellular components of epidermis and hair follicles (with the exception of hair follicle melanocytes) with strong functional activity. This finding would have clinical relevance to explain some of the cutaneous symptoms of Graves’ disease (Thiboutot, 1995). Thus, the observed expression of TSH-R on epidermal melanocytes may be connected with the skin pigmentation of Graves’disease as its intracellular mediator, cAMP acts as stimulator of melanocytes proliferation and differentiation. Indeed, both human and rodent malignant melanocytes and keratinocytes showed that TSH and anti-TSH-R antibodies stimulated cAMP production (Table III). Alternatively, expression of the TSH-R may turn melanocytes into targets for the destruction by TSH-R autoantibodies, thus explaining the high incidence of vitiligo in patients with Graves’ disease (Slominski et al, 1989). Also intriguing is the presence of TSH-R on hair follicle keratinocytes and papilla fibroblasts that might be involved in the association of Graves’ disease with alopecia areata. Finally, taken together with previous findings (Wu et al, 1996; Stadlmayr et al, 1997; Rapoport et al, 2000), the present observations raise the tantalizing prospect that some forms of Graves’ disease could be due to an autoimmune response directed primarily against a cutaneous TSH-R antigen. Thus, this could result from abnormal expression of major histocompatibility complex proteins, and abnormal exposure of TSH-R antigen to immune cells (Shimojo et al, 1996; Suzuki et al, 1999; Kohn et al, 2000), in response to exposure to environmental insults, e.g., solar radiation or skin infections.

In the thyroid, TSH-R-stimulated pathways regulate most aspects of intracellular iodide and thyroglobulin metabolism, synthesis of NIS and production, and secretion of T₄ (Kohn et al, 2000). We therefore evaluated the possibility of similar effects in the skin. Using reverse transcription–PCR we tested skin expression of TSH-R-regulated genes in vitro in stable cell lines positive for TSH-R (HaCaT keratinocytes and four different melanoma lines, e.g., SWM 35, WM164, WM1341D), and also in one line negative for TSH-R (SKMEL188). The lines expressing TSH-R also coexpressed thyroglobulin and NIS genes, whereas the SKMEL188 line was negative for these genes (Fig 1). All of these lines were negative for thyroid peroxidase gene (not shown). Thus, expression of some, but not all molecular elements of the TSH-R-related pathways that are characteristic for the thyroid is conserved in cultured skin cells.

Recent studies have shown NIS expression in lactating breast tissue and in some breast cancers (Spitzweg et al, 2001). TSH-R has been found on fat cells and other nonthyroid tissues (Szkudlinski et al, 2001). Functional roles have not been fully clarified for NIS in breast tissue or TSH-R in fat tissue, nor is it clear why such genes might be expressed in a pathologic situation. The existence of TSH-R in melanoma cells might be exploited if the TSH-R is involved in the regulation of growth or function of these cells as might be the case for NIS in breast cancer. Growth and function studies are in progress in our laboratories.

Reverse transcription–PCR assays for TSHβ, D2, D3, TRH-R, and TRH

We used the reverse transcription–PCR technique with sequencing of the amplified gene products together to define cutaneous expression of the genes coding for TSHβ, D2, D3, TRH-R, and TRH genes, in a combination of human tissue and cultured skin cells (Fig. 3; Table IV).

Figure 3. Expression of thyroid-related genes in human tissues.

(A) Expression of TSHβ: (1) DNA ladder; (2) pituitary; (3) follicular melanocytes; (4) SKMEL188 melanoma; (5) WM164 melanoma; (6) follicular keratinocytes; (7) epidermal keratinocytes; (8) dermal fibroblasts; (9) hair follicle papilla fibroblasts; (10) HaCaT keratinocytes. (B) Expression of D2 in human skin samples: (1,13) DNA ladder; (2) normal skin; (3) BCC; (4) HaCaT keratinocytes; (5) neonatal keratinocytes; (6) follicular keratinocytes; (7) epidermal keratinocytes; (8) neonatal keratinocytes; (9) dermal fibroblasts; (10) hair follicle papilla fibroblasts; (11) SKMEL188 melanoma; (12) SBCE2 melanoma. Arrows marked 227 bp (D2a) and 335 bp (D2b) transcripts. (C) Expression of D3 in human tissues and skin cells: (1,9) DNA ladder; (2) pituitary; (3,4) normal skin; (5) BCC; (6) myometrium; (7) follicular keratinocytes; (8) epidermal keratinocytes; (10) epidermal melanocytes; (11–13) SBCE2,WM35, and WM98 melanomas, respectively. (D) Expression of TRH-R: (1) DNA ladder; (2) pituitary; (3,4) SKMEL188 and WM1341D melanomas, respectively. (E) Expression of TRH: (1,10) DNA ladder; (2) nonlesional human skin; (3) dermal fibroblasts; (4) hair follicle papilla fibroblasts; (5) melanoma WM1341D; (6) neonatal keratinocytes; (7) adrenal gland; (8) myometrium; (9) placenta.

TSHβ

As we had found functional expression of TSH-R in the skin, we tested for its natural ligand (TSHβ) to investigate paracrine or autocrine control of TSH-R activity. Aside from its expression in the pituitary control, TSHβ gene expression was observed in epidermal and hair follicle keratinocytes, hair follicle melanocytes, squamous cell carcinoma, and two melanoma lines, as well as myometrium, being absent in all other cells or tissues tested that included biopsies of whole skin (Fig 3A; Table II). This pattern suggests that TSH expression is not constitutive, but instead responds to environmental stimulation exemplified, in this case, by the conditions of cell culture and/or by random depression during malignant progression.

D2 and D3 genes

The D2 gene product (227 bp) was expressed in almost all tested samples; the sole exception was a line of hair follicle melanocytes where it was below detectable levels (Fig 3B; Table II). In some biopsies of normal skin (one of four) and skin containing BCC (one of three), an additional second transcript of 335 bp was detected. It corresponds to the b isoform of D2 (D2b, accession no. AB041843). D2b contains an additional exon (insertion of 108 bp) with an in-frame TGA codon that may encode an extra selenocysteine (Ohba et al, 2001). This second isoform (335 bp) was also present in neonatal, epidermal, follicular and HaCaT keratinocytes, dermal and follicular papilla fibroblasts, epidermal melanocytes, squamous cell carcinoma cells, and the SBCE2 melanoma line (Fig 1B). The detection of the second isoform is consistent with the recent findings of two additional splicing variants of human D2 differentially expressed in brain, kidney, lung, and trachea (Ohba et al, 2001).

The D3 gene was expressed in all tissues tested that included brain, pituitary, adrenal gland, and intrauterine tissues. It was also detected in skin, where its expression was cell type specific (Fig 3C). It was expressed in all tested keratinocytes and fibroblasts lines, but was absent in melanocytes and in the majority of melanoma lines (five of six) (Table II).

D2 catalyzes 5′ (outer ring) deiodination of T₄ to T₃, whereas D3 catalyzes 5 (inner ring) deiodination of T₄ and T₃ to inactive T₃ (rT₃) and 3,3′-diiodothyronine (T₂) (St Germain, 1999; Bianco et al, 2002). The structure of both genes is brown. D2 is predominantly expressed in pituitary, brain, brown fat, thyroid, heart, and skeletal muscle, whereas D3 is predominantly expressed in the brain, uterus, placenta, fetal membranes, and skin (rodents) (St Germain, 1999). Actual transformation of T₄ to T₃, and of T₃ to T₂ have been demonstrated in cultured epidermal keratinocytes (Kaplan et al, 1988), although the pattern of expression of the corresponding genes has not been evaluated in human skin. Our data thus represent the first demonstration of D2 and D3 gene expression in human skin. A differential pattern for expression of the genes becomes clearly evident along the predominant cellular components of the epidermis and dermis. The absence of D2 in hair follicle melanocytes and of D3 in both epidermal and hair follicle melanocytes, in the majority of melanoma cells and in the squamous cell carcinoma line suggest cell lineage specific gene expression and probably, of correspondingly specific patterns of T3 degradation in the same populations. Future biochemical studies on the differential pattern of cell type specific T3 production and degradation should clarify the functional implications at the systemic level, and in epidermal and dermal homeostasis and activity of adnexal structures.

TRH and TRH-R

As TSH production is under the control of hypothalamic TRH, we evaluated local expression of the TRH-R gene. As expected, expression of TRH-R gene was found in brain and pituitary. In the skin, it was detected only in a single skin biopsy specimen of BCC, and in two melanoma lines SKMEL188 and WM1341D (Fig 3D; Table II). Thus, TRH-R may not participate directly in the physiology of the skin. Conversely, TRH that was detected in brain, adrenal gland, myometrium, placenta, umbilical cord, and fetal membrane, was also widely present in skin cells. It was found in normal skin (in one of four biopsies), in four melanoma lines, in neonatal keratinocytes and in both dermal and follicular papilla fibroblasts (Fig 3E, Table II). Therefore, TRH is not coexpressed with TRH-R in most cells (with exception of SKMEL188 melanoma) suggesting that paracrine or autocrine mechanisms may not operate locally in peripheral tissues, e.g., skin, placenta, fetal membranes, uterus, and adrenal gland. It can be speculated that expression of TRH mRNA by the above tissues and cell types may result in TRH production mainly for export, to regulate regional (skin fibroblasts, adrenal gland) or fetal (placenta, umbilical cord, fetal membranes) homeostasis. Of note, amphibian skin regulates pituitary function through local production of TRH that is exported to stimulate pituitary production and release of prolactin and α-melanoctye-stimulating hormone (Vaudry et al, 1999). Thus, this peptide gene expression may very well represent an evolutionary continuum.

CONCLUSIONS

There is broad cutaneous expression of the gene and functional TSH-R, which is still of undetermined physiologic and pathologic significance. Genes for elements of the hypothalamic–pituitary–thyroid axis are also expressed in the skin, but with selectivity in both cell type and gene type. It appears that there may be important divergences between the roles of the molecules at the central and peripheral levels.