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Functionally Impaired TR Mutants Are Present in Thyroid Papillary Cancer

Department of Endocrinology, Medical Research Center, Polish Academy of Sciences (M.P.-K., A.K., A.M., J.N.), and Department of Biochemistry, Medical Center of Postgraduate Education (M.P.-K.), 02-097 Warsaw, Poland; and Laboratory of Molecular Biology, National Cancer Institute, National Institutes of Health (S.-Y.C.), Bethesda, Maryland 20892

Address all correspondence and requests for reprints to: Dr. Monika Puzianowska-Kuznicka, Department of Endocrinology, Medical Research Center, 1a Banacha Street, 02-097 Warsaw, Poland. E-mail: . monika{at}amwaw.edu.pl

Abstract

TRs are transcription factors that regulate cell proliferation, differentiation, and apoptosis. They are cellular homologs of the transcriptionally inactive viral oncogene v-erbA. We tested the hypothesis that the functions of TRs could be impaired in cancer tissues as a result of aberrant expression and/or somatic mutations. As a model system, we selected human thyroid papillary cancer, in which the most common abnormalities, RET/papillary thyroid cancer rearrangements (fusion of RET kinase domain to the activating domains of other genes), were found in 40–45% of cases. We found that the mean expression levels of TRß mRNA and TR mRNA were significantly lower, whereas the protein levels of TRß1 and TR1 were higher in cancer tissues than in healthy thyroid. Sequencing of TRß1 and TR1 cDNAs, cloned from 16 papillary cancers, revealed that mutations affected receptor amino acid sequences in 93.75% and 62.5% of cases, respectively. In contrast, no mutations were found in healthy thyroid controls, and only 11.11% and 22.22% of thyroid adenomas had such TRß1 or TR1 mutations, respectively. The majority of the mutated TRs lost their trans-activation function and exhibited dominant negative activity. These findings suggest a possible role for mutated thyroid hormone receptors in the tumorigenesis of human papillary thyroid carcinoma.

TUMORIGENESIS IS a complex, multistep process, involving many oncogenes and/or tumor suppressors that are either considered specific for a certain tumor type [e.g. VHL (von Hippel-Lindau) in renal clear cell cancer (1), BRCA1 (breast cancer 1) in breast and ovarian cancer (2)] or involved in the tumorigenesis of many different cancers (e.g. p53). Nevertheless, the process of tumorigenesis is not completely understood, and the function and identity of only a limited number of factors involved in this process are known. Nuclear TRs are good candidates for being such factors. TR1 is a cellular homolog of the viral v-erbA oncogene, a transcriptionally inactive, dominant negative receptor (3, 4) that has been shown to be involved in the neogenesis of avian erythroblastosis, some sarcomas (5, 6), liver cancer, and thyroid abnormalities (7). In contrast to v-erbA, transcription activation of downstream genes by TRs is necessary for proper tissue and developmental stage-dependent regulation of proliferation, differentiation, and apoptosis (8). TRs mediate these diverse biological effects by heterodimers formed with other nuclear proteins (8, 9); by their ability to bind DNA as monomers, homo-, and heterodimers (10, 11); and by their ability to recognize different DNA sequences (direct repeats, DR4; palindromes, P0; inverted palindromes, Ip6) (12, 13). The function of TRs is modulated by the interactions with many proteins that are ubiquitously distributed or expressed in a tissue- or a cell-specific manner (14, 15, 16, 17). The function of TRs can be significantly affected by the alteration of their genes. It has been shown that LOH of the 3p21-p25 region, where the TRB gene is located (18, 19), was observed in up to 100% of small cell lung cancers (19, 20), 60% of uveal melanomas (21), and 64% of nonfamilial renal cell carcinomas (22), whereas LOH of the 17q21 region, where the TRA gene is located (23), was found in 79% of breast cancers (24). A high incidence of TR mutations was detected in hepatocellular tumors (25). TRs mRNA and/or protein amounts were found to be significantly changed in different cancers (25, 26, 27, 28). In the present study we chose human papillary thyroid cancer (PTC) as a model system to understand the involvement of TRs in tumorigenesis. Interestingly, the available data show that common tumor suppressors and oncogenes are not significantly affected in this type of cancer. For example, no or only a low frequency of abnormalities of P53 (29, 30, 31) and retinoblastoma (RB) tumor suppressors (32) or ERB-2 (33), RAS (30, 31), and FHIT (34) oncogenes were observed in this type of malignancy. In fact, only RET/PTC rearrangements (fusion of RET tyrosine kinase domain to activating domains of other factors, considered specific to PTC) were detected in 40–45% of the analyzed cases (31, 33, 35). Here we show that TRs in PTC are affected by the aberrant expression at both the mRNA and protein levels and by a high incidence of mutations resulting in alteration of protein sequence and functions. We propose that functionally impaired TR play an important role in tumorigenesis.

Materials and Methods

Tissues

Tissues were obtained (with the permission of the committee of human investigation) during total thyroidectomies performed due to the diagnosis of the cancer. A fragment of the tumor (22 carcinomas, as checked visually during surgery and confirmed by histopathological evaluation of tumor borders) and part of the opposite thyroid lobe not infiltrated with cancer (22 healthy controls) were excised, immediately frozen on dry ice, and stored at -75 C until needed. The final diagnosis of PTC was established upon histological evaluation of the excised organs. Similarly, 10 follicular adenomas were collected as hypertrophic nonmalignant controls.

RNA isolation

Twenty to 100 mg deep frozen tissue were homogenized in a glass-Teflon homogenizer directly in 1 ml TRIzol reagent (Life Technologies, Inc., Gaithersburg, MD). Samples were incubated at room temperature for 5–10 min, supplemented with 0.2 ml chloroform, mixed by shaking for 15 sec, incubated at room temperature for 10 min, then centrifuged at 12,000 x g for 15 min at 4 C. To precipitate RNA, the upper, aqueous phase was transferred to another Eppendorf tube, mixed with 0.5 ml isopropyl alcohol, incubated at room temperature for 15 min, and centrifuged at 12,000 x g for 15 min at 4 C. The RNA pellet was washed with 75% ethanol, briefly dried, and resuspended in 40 µl diethylpyrocarbonate-treated water. The RNA concentration was calculated from spectrophotometry measurements.

Northern blot hybridization

Total RNA (2.5 µg) was electrophoresed on 1.2% agarose-formaldehyde gel, partially hydrolyzed by NaOH treatment, and transferred onto a GeneScreen membrane (NEN Life Science Products, Boston, MA). To check for RNA loading and quality and to visualize 18/28S rRNA, Northern blots were stained with methylene blue, photographed, and destained before hybridization. Hybridization was performed with probes made of full-length coding regions of TR1 (1232 bp) or TRß1 (1370 bp). Probes were prepared with Megaprime DNA Labeling System (Amersham Pharmacia Biotech, Madison, WI) by a random priming method in the presence of [-³²P]dCTP (SA, 6000 Ci/mmol; NEN Life Science Products, Little Chalfont, UK). After overnight hybridization at 42 C (50% formamide, 5 x saline-sodium phosphate EDTA, 0.2% SDS, 10% dextran sulfate, 5 x Denhardt’s solution, and 100 µg/ml denatured sheared salmon sperm DNA), the filters were washed three times for 5 min each time at room temperature in 2 x SSC/0.2% SDS. Stringent washes were then performed twice for 15–20 min each time in 0.25 x SSC/0.1% SDS at 55 C. Hybridized blots were exposed against Kodak film (NEN Life Science Products) for 1–6 d. The intensities of specific bands were measured by densitometry, and the results were normalized against the 18/28S bands of rRNA and for the purpose of statistical analysis expressed in arbitrary units.

Isolation of nuclear proteins

The buffers used for isolation were described by Kane et al. (36). All buffers were supplemented with pepstatin A (final concentration, 1 µg/ml), leupeptin (1 µg/ml), aprotinin (2 µg/ml), and phenylmethylsulfonylfluoride (0.5 mM). Twenty to 100 mg tissue were homogenized in a glass-Teflon homogenizer in 1 ml ice-cold STM buffer (0.25 M sucrose, 20 mM Tris-HCl, and 1.1 mM MgCl₂, pH 7.85) and centrifuged at 1,000 x g for 10 min at 4 C. The pellet was washed twice in 1 ml STM buffer with 0.5% Triton X-100, then centrifuged under the conditions described above. The final pellet was resuspended in 0.1 ml STM with potassium KSTM and 20% glycerol buffer (0.25 M sucrose, 20 mM Tris HCl, 1.1 mM MgCl₂, 0.4 M KCl, 20% glycerol, 5 mM dithiothreitol), sonicated, and incubated on ice for 30 min with vortexing every 5 min to extract the soluble nuclear proteins. After solubilization, the suspension was centrifuged at 12,000x g for 15 min at 4 C. The amount of protein in the supernatant was quantified by spectrophotometry with Bradford’s reagent (protein assay, Bio-Rad Laboratories, Inc., Munich, Germany). Proteins were stored in 10-µl aliquots at -75 C.

Western blots

Twenty micrograms of each nuclear extract were resolved by SDS-PAGE (10% gel). Proteins were transferred onto a nitrocellulose membrane (Bio-Rad Laboratories, Inc.). Protein loading was additionally monitored by Ponceau Red staining. Destained membranes were blocked overnight at 4 C in PBS and 0.1% Tween 20 (PBS-T buffer) supplemented with 5% nonfat dry milk powder, washed twice in PBS-T for 10 min at room temperature, then incubated with rabbit polyclonal anti-TR1 primary antibody (1:1000 in PBS-T; Affinity BioReagents, Inc., Golden, CO) or mouse monoclonal anti-TRß1 primary antibody (1:1000; Dr. S.-Y. Cheng, NIH, Bethesda, MD) for 1 h, washed three times for 10 min each time in PBS-T, incubated for 1 h at room temperature with the secondary antibody (horseradish peroxidase-conjugated antirabbit or antimouse, respectively), and washed again five times for 10 min each time in PBS-T. Specific receptor proteins were visualized by chemiluminescent reaction performed with an ECL kit (Amersham Pharmacia Biotech, Little Chalfont, UK). Exposure to Kodak film was for 15 sec to 15 min. After stripping, blots were incubated with mouse monoclonal anti-ß-actin antibodies (1:5000 in PBS-T; Sigma, St. Louis, MO) and processed as described above. The intensities of specific bands were measured by densitometry, and the results were normalized against ß-actin and, for the purpose of statistical analysis, expressed in arbitrary units.

Gel retardation assays (electrophorectic mobility shift assay)

The assays were performed with 7.5–10 µg of the nuclear extracts isolated from tumors and their respective controls. The probe was a double-stranded DNA containing thyroid response element (TRE; 5'-GATCGCAGGTCATTTCAGGACAGCGAT-3'; TRE-DR4). Mutated TRE served as a nonspecific competitor (5'-GGCAAATCATTTCAAGACAG-3'). Nuclear extracts were incubated at room temperature for 20 min in a binding buffer [20 mM Tris-HCl (pH 7.5), 50 mM KCl, 2 mM dithiothreitol, 0.1% Triton X-100, and 6% glycerol] with 250 ng dI-dC, 1 ng probe, and a 10-fold excess of the specific or nonspecific competitor. In supershift experiments 1 µl of above-described anti-TR1 or anti-TRß1 antibody was added to the binding reaction (instead of competitor), and incubation was performed on ice for 30 min and subsequently at room temperature for 20 min. Products of the reaction were loaded onto a 4% native gel and electrophoresed at 150 V for 2 h at room temperature, then the gel was dried and exposed against Kodak film for 1–2 d.

Cloning of TR1 and TRß1 cDNA from PTC and control tissues

To clone TR1 or TRß1 expressed in PTC, RT of 1 µg total RNA was performed with Superscript II reverse transcriptase (Life Technologies, Inc.) and T₁₈ primer. One tenth of the total volume of this reaction was used as a template in a regular PCR reaction, performed with increased fidelity ExTaq polymerase (TaKaRa Shuzo, Otsu, Japan). To clone TR1 the following primers were used: forward, 5'-GGATGGAATTGTGAATG-3' or 5'-ATGGAACAGAAGCCAAGCAA-3'; and reverse, 5'-GGCCGCCTGAGGCTTTA-3'. To clone TRß1 the following primers were used in the PCR reaction: forward, 5'-GATCCAGAATGATTACTAACC-3'; and reverse, 5'-GGAATTATAGGAAGGAATCC-3'. The cDNA amplification was performed as follows: 3 min at 94 C, then 35–40 cycles of 94 C for 40 sec, 52 C for 1 min, 72 C for 5 min, and a 10-min final extension at 72 C. As the amount of TRß1 cDNA was very low after the first PCR, the product of this PCR was electrophoresed on 1% agarose gel, the gel fragment of the expected TRß1 size was excised, dissolved in 100 µl distilled deionized water, and 5 µl of this solution were used as a template in a second PCR, that was performed with the following internal primers: forward, 5'-ATGACAGAAAATGGCCTTAC-3'; and reverse, 5'-CTAATCCTCGAACACTTCCA-3'. The reaction conditions were as described above, except that only 25 cycles were performed. The final PCR products were electrophoresed on 1% agarose gel, specific TR1 and TRß1 bands were excised, DNA was isolated from the gel with a QIAquick Gel Extraction Kit (QIAGEN, Hilden, Germany) and ligated into pGEM-T (TR1) or pGEM-T Easy (TRß1) vector containing T overhangs (Promega Corp., Madison, WI) at 4 C (overnight incubation). JM109 bacteria were transformed with ligation mix, and blue-white selection was performed. Mini-preps were performed using the Wizard Plus SV Minipreps DNA Purification System (Promega Corp.) or Plasmid Miniprep Plus (A&A Biotechnology, Gdynia, Poland) kits. TR1 and TRß1 clones underwent restriction analysis (TR1 with NcoI and TRß1 with BstXI) to establish the insert position within the pGEM-T vector.

Sequencing of the TR clones

Automatic sequencing of TR1 or TRß1 clones was performed with the BigDye Terminator Cycle Sequencing Kit (Perkin-Elmer Corp., Wellesley, MA). At least 3 TR1 or 3 TRß1 clones originating from the same cancer tissue were pooled and sequenced. Analysis was performed with Lasergene software (DNASTAR, Madison, WI). Upon mutation detection, individual clones were then manually resequenced with the T7 Sequenase version 2.0 DNA sequencing kit (U.S. Biochemical Corp., Cleveland, OH). In addition, TR1 and TRß1 from 9 thyroid adenomas as well as 11 and 16 (for TR1 and TRß1, respectively) healthy thyroid tissues originating from thyroid lobes opposite to PTCs with confirmed mutations were sequenced.

Cloning of the mutated TR1 and TRß1 into pcDNA3.1(+) eukaryotic expression vector

pGEM-T constructs containing TR1 mutants positioned with their 5'-ends next to the T7 promoter were cut with ApaI, treated with Klenow enzyme to blunt ends, then restricted with NotI and recloned into pcDNA3.1(+) vector (Invitrogen, Carlsbad, CA) previously prepared with EcoRV and NotI. TR1 mutants cloned in the opposite direction in pGEM-T (with their 5'-ends next to the SP6 promoter) were cut with ApaI and NotI enzymes and recloned into pcDNA3.1(+) vector prepared with the same enzymes. TRß1 mutants cloned into pGEM-T Easy vector with their 5'-ends next to T7 promoter were cut with NcoI, treated with Klenow enzyme, then restricted with NotI and recloned into pcDNA3.1(+) vector cut with EcoRV and NotI. TRß1 mutants cloned in the opposite direction within the pGEM-T Easy vector (with their 5'-ends next to SP6 promoter) were restricted with SpeI, treated with Klenow, restricted with NcoI, and recloned into pcDNA3.1(+) cut with EcoRV and NotI. To confirm the position of recloned mutants within final constructs, restriction analysis was performed with BstXI.

Transcription activation by TR1 and TRß1 mutants

pGL2-Promoter vector (Promega Corp.) containing firefly luciferase gene driven by the simian virus 40 (SV40) promoter served as reporter vector in the transcription activation tests. In front of the promoter, a human 5'-deiodinase type I TRE (5'-GGGACTAGTAGGCTATCTGAGGTCAGGAGTTCAAGA3') was cloned (pGL2-TRE vector). A pRL-cytomegalovirus (CMV) vector (Promega Corp.) containing the Renilla luciferase gene served as an internal control, and all results were normalized against this control. A triple transfection of HEK 293 cells was performed with 0.2 µg pGL2-TRE, 0.2 µg pRL-CMV, and 0.2 µg pcDNA3.1(+)-mTR (mutant) or pcDNA3.1(+)-wtTR (wild-type). The reaction was performed in 12- or 24-well plates containing the same number of plated cells, with Lipofectamine Plus reagent (Life Technologies, Inc.). Transfection was performed according to the manufacturer’s suggestions in the medium with no serum with the following modifications: in total, 0.6–0.8 µg DNA, 1–2 µl Lipofectamine, and 4 µl Plus reagent were used. The incubation lasted for up to 2 h, then medium, FCS (to a final concentration of 10%), and T₃ (to a final concentration of 100 nM) were added. The incubation lasted for 24 h before lysis of the cells. The luciferase assay was performed with the Dual Luciferase Reporter Assay System (Promega Corp.), strictly following the protocol supplied by the manufacturer, and measurements were made with a Turner Designs luminometer (Turner Designs, Sunnyvale, CA). Each experiment was repeated five times.

Dominant negative function of TR1 and TRß1 mutants

All vectors and procedures were as described above, except that in addition to pGL2-TRE and pRL-CMV vectors, both pcDNA3.1(+)-mTR and pcDNA3.1(+)-wtTR were cotransfected, respectively, at 1:1 and 3:1 ratios.

Results

Expression of TRß and TR mRNA in PTC

Total RNA was isolated from three types of tissues: PTC, the opposite lobe of thyroid with no signs of tumor infiltration, and adenoma (follicular) nodules. Altogether RNAs from 22 tumors, 22 healthy controls from the same thyroid, and 10 adenomas were analyzed by Northern blot. The human TRß1 cDNA was used as a TRß probe for hybridization. In 95.45% of the analyzed PTCs, the amount of TRß mRNA was lower in the cancer than in its respective control. In only 1 case (4.55%) was the opposite situation observed (Fig. 1B). The maximum difference in TRß mRNA levels found in tumor and healthy tissue, both excised from the same thyroid, was 27-fold (single case). Four specific bands, a dominant 10-kb band and much less prominent bands of approximately 5.0, 3.0, and 2.0 kb (37), were consistently present in all examined samples (Fig. 1A). The mean amount of TRß mRNA expressed in arbitrary units was 0.51 ± 0.24 in PTC, 1.79 ± 0.9 in healthy thyroid, and 1.84 ± 0.11 in adenomas. Overall, the amount of specific mRNA was 3.5 times lower in PTC than in healthy tissue (by Mann-Whitney U test, P < 0.001) and 3.6 times lower than in adenoma nodules (by Mann-Whitney U test, P < 0.001). Hybridization with a TR probe corresponding to the open reading frame of TR1 revealed the presence of 2 bands of approximately 6.0 and 3.2 kb (Fig. 1C). In 86.35% of the analyzed PTCs, the amount of TR mRNA was lower, in 4.55% it was almost identical, and in 9.1% it was higher in tumor tissue than in healthy control tissue (Fig. 1D). The maximum difference in TR expression levels between the tumor and its respective healthy control was 4.17-fold. The mean amount of TR mRNA expressed in arbitrary units was 0.46 ± 0.25 in PTC, 0.82 ± 0.35 in healthy thyroid, and 0.98 ± 0.06 in adenoma. The mean amount of this mRNA in cancer tissues was 1.77 times lower than that in healthy controls (by Mann-Whitney U test, P < 0.001) and approximately 2 times lower than in adenomas (by Mann-Whitney U test, P < 0.001). When the upper 6.0-kb band (representing TR1 mRNA only) and the lower 3.2-kb band (representing TR1 and TR2 mRNA) (37) were evaluated separately, the same tendency was observed; their intensities were 1.71 and 1.78 times lower, respectively, than that of healthy thyroid and 2.12 and 2.0 times lower, respectively, than that of adenoma nodules.

View larger version (38K): [in this window] [in a new window]   Figure 1. Northern blots of total RNA isolated from PTC and healthy, opposite thyroid lobes. Samples (2.5 µg) of total RNA were resolved on 1.2% agarose-formaldehyde gels and blotted onto a positively charged nylon membrane. Hybridizations were performed with the probes made of the TRß1 and TR1 cDNAs. The intensity of specific mRNA bands was normalized against 28S and 18S ribosomal RNA. CA, PTC; C, control, healthy opposite thyroid lobe. A, Representative Northern blots hybridized with TRß1 probe. 10, 8, 6, Numbers of cancer cases. B, TRß mRNA amount (arbitrary units) in all analyzed cancer/control pairs. C, Representative Northern blots hybridized with TR1 probe. 19, 20, 8, Numbers of cancer cases. D, TR mRNA amount (arbitrary units) in all analyzed cancer/control pairs.

Expression of TR1 and TRß1 proteins in PTC

To evaluate the protein expression levels of TRs, Western blot analysis of nuclear extracts isolated from PTC tumors and their respective controls was performed (Fig. 2, A and C). Due to the limited amount of PTC tissues, only 18 tumor/control pairs were analyzed. In contrast to the mRNA results, in 88.88% cases an increased level of TRß1 protein was found in tumor tissues (ranging from 1.1- to 6.4-fold), whereas in remaining 11.12% this level was decreased compared with that in the respective controls (Fig. 2B). The mean amount of TRß1 protein expressed in arbitrary units was 6.74 ± 4.02 in PTC, 3.62 ± 2.10 in healthy thyroid, and 5.62 ± 1.32 in adenomas. In general, the mean amount of this protein was 1.86-fold higher in tumor tissues than in healthy controls (by Mann-Whitney U test, P < 0.005) and 1.27-fold higher than in adenomas (statistically not significant). The ratio of TRß1 protein amount in a given tumor and its respective control was not proportional to the ratio of TRß mRNA amount in the same tissues. The level of TR1 protein was increased in 72.22% and decreased in 27.78% of the analyzed tumors compared with their respective controls (Fig. 2D). The mean amount of TR1 protein expressed in arbitrary units was 5.51 ± 3.80 in PTC, 4.05 ± 2.42 in healthy thyroid, and 3.20 ± 1.05 in adenoma. The overall mean amount of TR1 protein was 1.36-fold higher in tumor tissues (statistically not significant) and 1.72-fold higher than in adenomas (by Mann-Whitney U test, P < 0.005). Similarly, there was no correlation between the ratio of TR mRNA amounts and TR1 protein amounts in tumor/control pairs.

View larger version (34K): [in this window] [in a new window]   Figure 2. Immunoblots of nuclear extracts isolated from PTC and healthy, opposite thyroid lobes. Samples (20 µg) of nuclear extracts were subjected to SDS-PAGE (10% gel) and transferred onto nitrocellulose membranes. Blots were incubated with primary monoclonal anti-TRß1 or primary polyclonal anti-TR1 and secondary horseradish-labeled antimouse or antirabbit (respectively) antibodies as well as with primary monoclonal anti-ß-actin, and secondary antimouse antibodies. A chemiluminescent reaction was performed with ECL reagents (Amersham Pharmacia Biotech). The intensity of specific protein bands was normalized against the intensity of the ß-actin band. CA, PTC; C, control, healthy opposite thyroid lobe. A, Representative immunoblots hybridized with anti-TRß1 and anti-ß-actin antibodies. 1, 2, 4, Numbers of cancer cases. B, TRß1 protein amount (arbitrary units) in all analyzed cancer/control pairs. C, Representative immunoblots hybridized with anti-TR1 and anti-ß-actin antibodies. 6, 9, 14, Numbers of cancer cases. D, TR1 protein amount (arbitrary units) in all analyzed cancer/control pairs.

To evaluate the ability of TRs present in cancer tissues to specifically bind DNA, gel retardation assays were performed with DNA containing the TRE-binding site. Nuclear extracts isolated from 16 PTC/control tissue pairs were available for this analysis. In 68.75% of the analyzed cases, the DNA binding by all TRs present in the cancer nuclear extracts was weaker, whereas in the remaining 31.25% it was stronger than by TRs present in nuclear extracts of their respective controls. In addition, in some cases a lack of certain bands was observed. No correlation of the amount of TR proteins (as checked by immunoblots) and the strength of DNA binding was observed, strongly suggesting the presence of TR mutations altering the DNA binding activity. In addition, supershift experiments were performed. The presence of anti-TR1 or anti-TRß1 antibodies in the reaction caused the appearance of more retarded bands, confirming that the observed shifts were due to the interactions of TRs present in nuclear extracts with the probe (Fig. 3).

View larger version (76K): [in this window] [in a new window]   Figure 3. Gel retardation assay (electrophorectic mobility shift assay) of representative papillary thyroid cancer and control, healthy thyroid nuclear extracts. Samples (7.5–10 µg) of nuclear extracts were incubated with the DNA probe (TRE), 250 ng dI-dC, specific competitor, or anti-TR1 or anti-TRß1 antibodies in the buffer containing 50 mM KCl. Products of the reaction were resolved on 4% native gels. CA, PTC; C, control, healthy opposite thyroid lobe; -, no competitor; 10x S, 10-fold molar excess of specific competitor; 1 AB, anti-TR1 antibodies; ß1 AB, anti-TRß1 antibodies; 3, number of the representative cancer case with weaker DNA binding.

Mutations of TR1 and TRß1 in PTC

RNAs from 16 tumors were available for the cloning of TR1 and TRß1. cDNAs were first cloned by the RT-PCR method and subsequently cloned into pGEM-T (TR1) or pGEM-T Easy (TRß1) vectors. The DNA sequence indicated that 100% of analyzed PTCs contained mutated TRß1. Altogether 44 TRß1 mutations were found, 14 of them silent (no amino acid substitution). In 93.75% of the analyzed PTCs, mutations resulted in amino acid substitutions. Two mutations present in TRß1 cloned from a single PTC (6.25%) were silent, resulting in a wild-type protein. In 43.75% of PTCs, TRß1 mutations were heterogeneous (mutated as well as wild-type cDNAs were found). Only 12.5% of TRß1 mutants had the same residue mutated (M32V, M32T). The number of mutations within a single clone (representing 1 allele) ranged from 1–5 (Table 1). Some 68.75% of PTCs had TR1 mutated. Altogether 30 mutations were found, 7 of them silent. In 62.5% of PTC cases mutations resulted in amino acid substitution. In 1 case (6.25%) a single silent mutation was present, resulting in a wild-type receptor protein. In 54.54% of PTCs where TR1 mutants were identified, mutations were heterogeneous. Interestingly, in 27.27% of such PTCs identified, TR1 mutants had the same residues mutated (S183N, H184Q, R228H). The number of mutations within a single clone (representing one allele) ranged from 1–6 (Table 1). Analysis of the predicted amino acid sequence of TR1 and TRß1 showed that amino acid substitutions resulting from gene mutations were distributed throughout the length of the protein. There were two points of concentration of TRß1 mutations, the first on the border of the A/B and DNA-binding domains and the second at the end of the ligand-binding domain, but no true "hot spots" were found. For control, TRs cloned from 9 thyroid adenomas as well as 16 and 11 healthy thyroid tissues (taken from thyroid lobes opposite PTCs with confirmed TRß₁ or TR₁ mutations, respectively) were sequenced. Sequencing of TRs cloned from thyroid adenomas revealed that TRß1 mutations were present in 44.44% of the cases. In 33.33% of the cases, mutations were silent (in 2 cases single, in one case 3 such substitutions), whereas in only 11.11% (one case) were 2 mutations affecting protein amino acid sequence (E390G, R391G) as well as 1 silent substitution present. Some 55.55% of the analyzed adenomas were TRß1 mutation free. TR1 mutations were found in 55.55% of the adenomas. In 33.33% of the cases, they were single silent mutations, in 11.11% (1 case) the mutation resulted in amino acid substitution (Y435H), and in 11.11% 1 mutation affecting protein amino acid sequence (I299T) as well as 1 silent substitution were present. Some 44.44% of the analyzed adenomas were free of TR1 mutations. Sequencing of TRs cloned from 16 and 11 healthy thyroid controls originating from thyroid lobes opposite PTCs with confirmed TRß1 or TR1 mutations, respectively, revealed that all were wild-type receptors.

View this table: [in this window] [in a new window]   Table 1. Predicted amino acid substitutions caused by somatic mutations of TR1 and TRß1 cDNA cloned from PTC

To assess the trans-activation activity of mutant TRs, a reporter construct pGL2-TRE containing firefly luciferase gene driven by SV40 promoter and TRE enhancer was activated by transfected TR proteins in the presence of 100 nM T₃. It was found that the reporter gene present in the original pGL2 vector was strongly activated by endogenous transcription factors activating the SV40 promoter. In addition, introduction of TRE into the reporter vector caused an approximately 2-fold increase in its transcription by the endogenous TRs. After deduction of background activation, it was found that 2 mutants containing silent mutations within their cDNAs (8 1 and 15 ß1) activated transcription similarly to the level of their respective wild-type receptors. All but 1 (3 ß1) TRß1 mutants with amino acid substitutions were defective in transcription activation. Two of them, both truncated mutants 6 (N76D, S81L, I135V, Q136H, R201X) and 9 (Q235X, M379T, D427G), were practically nonfunctional (Fig. 4A). Similarly, when compared with wild-type TR1, all TR1 mutants containing amino acid substitutions presented weakened trans-activation activity. Two truncated mutants, highly mutated 13 (S183N, H184Q, Q187X, R228H, E245V, K288E) and 11 (K29T, C97X), were transcriptionally inactive, whereas the most active mutant 9 (G57E) retained 70.87% of wild-type TR1 activity (Fig. 4B).

View larger version (27K): [in this window] [in a new window]   Figure 4. The trans-activation activity of mutant TR cloned from PTC. The expression pcDNA3.1(+) plasmids of wild-type or mutant thyroid hormone receptors ß1 (A) and 1 (B) were cotransfected with reporter pGL2-TRE into HEK293 cells. After 24-h incubation with 100 nM T3, cell lysates were prepared and reporter firefly luciferase activity was determined and normalized against the activity of control Renilla luciferase. The trans-activation activity of analyzed mutants is shown in relation to wild-type receptor activity (100%; mean ± SD; n = 5).

To determine whether the TR mutants can act as dominant negative receptors, HEK293 cells were cotransfected with mutant and wild-type TR-encoding constructs at ratios of 1:1, and 3:1, respectively. Cells were incubated for 24 h in the presence of 100 nM T₃. After incubation, luciferase activity was assayed as described above. At a 1:1 ratio, there were three mutants (11 1, 3 ß1, 14 ß1) with no or almost no dominant negative activity, while mutant 9 ß1 had the highest such activity. The dominant negative activity of most mutants ranged from 40–70%. Although in the majority of analyzed cases a 3-fold excess of mutant TR intensified its dominant negative effect on its respective wild-type TR (e.g. 2 1 and 14 ß1), in some cases this effect remained similar to that observed at a 1:1 mutant to wild-type TR ratio (e.g. 9 1 and 13 ß1) (Table 2).

View this table: [in this window] [in a new window]   Table 2. Dominant-negative potency of TR1 and TRß1 mutants (at 100 nM T3; % of wild-type receptor activity) cloned form PTC

By analogy to the v-erbA oncogene, we postulated that TRs present in cancer tissues could be mutated and transcriptionally impaired. Indeed, in this paper we describe the highest incidence of genetic alterations affecting a factor potentially involved in the tumorigenesis of papillary thyroid cancer. Until now, the most common abnormalities described in this cancer were RET/PTC rearrangements, found in up to 45% of cases (31, 33). The present work shows that the frequency of PTC cases affected by TRß1 or TR1 mutations resulting in amino acid substitutions is much higher, reaching 93.75% and 62.5%, respectively. It is interesting that the number of such mutations found in a single TRß1 or TR1 allele was two to six in approximately 60% and 40% of all analyzed PTC cases, respectively. This finding suggests the presence of a potential DNA repair defect in cancer tissues and that extensive mutation and a marked impairment of the receptor function have to occur for TRs to influence the process of tumorigenesis. It is difficult to establish, however, which mutation and to what extent leads to the loss of a specific function of the receptor (such as DNA binding, T₃ binding, heterodimerization with RXR, etc.), especially that the contributions of these mutations may not be additive (38). The frequency of TR mutations resulting in amino acid substitutions is low in nonmalignant thyroid hypertrophic changes, i.e. adenomas (11.11% and 22.22% for TRß1 and TR1, respectively). No TRß1 or TR1 mutations were present in healthy thyroid controls, confirming that mutations found in PTCs were somatic. As expected, we found that all but one mutant (96%) were transcriptionally impaired (mostly from 2- 50% of the retained wild-type activity). In addition, all but three mutants (88%) exhibited a dominant negative activity.

We extended our observations to other levels of TR regulation control. We showed that the mean decrease in specific TRß mRNA amount in PTC compared with the controls was significant (3.5-fold), as it was for TR mRNA (1.77-fold). Interestingly, in contrast to the results obtained at the mRNA level, the mean amounts of TRß1 and TR1 proteins (two major transcriptionally active T₃ receptors) were higher in PTC than in controls. It is important to note that the decreased mean amounts of TR mRNA and the increased mean amounts of TRß1 and TR1 proteins were associated only with malignant tumors, not with adenomas. We also observed impaired DNA binding by TRs in tumor tissues. Based on these observations, we suggest that the transcription of TR downstream genes that are responsible for the regulation of cell differentiation, apoptosis, and proliferation could be markedly altered in tumor tissues. In such a situation the cell phenotype could be changed to that of a less differentiated cell that again could respond to proliferative signals. In addition, cells with DNA defects would not be eliminated by means of apoptosis. Our hypothesis seems to be supported by the findings of other researchers. For example, the sodium-iodide symporter gene (NIS) that contains a TRE within the 5'-flanking region (39) is down-regulated in thyroid cancers (40), possibly due to poor activation of downstream genes by mutated TRs. It has been shown that overexpression of wild-type TRß1 protein caused a significant increase in the expression of TSH receptor in thyroid cells (41). Therefore, the altered function of TRß1 protein described in this paper could be in part responsible for the decreased TSH receptor expression observed in PTC (42). The expression of other thyroid differentiation markers is associated with the function of TRs via the nuclear factor I (NFI) family of transcription factors. NFI genes have been shown to be T₃ response genes in the Xenopus laevis organogenesis model (43). Their human homologs bind to their recognition sites within the promoters of thyroid transcription factor 1 (44) and thyroid peroxidase (45) genes, enhancing their expression. Therefore, the low expression of thyroid transcription factor 1 (46) and thyroid peroxidase (40) genes observed in PTC could in part be an indirect result of disturbed activation of NFI genes by mutated TRs. Interestingly, a negative T₃ response element has been identified in the promoter of the c-myc gene, a factor that is required for cell cycle progression (47). The altered binding or the binding of mutated TR with impaired T₃-binding activity to negative TRE could then activate the expression of c-myc. Indeed, the expression of this factor in PTC is increased (48, 49) and has been shown to correlate with a low amount of TRß mRNA (42). It has also been shown that unliganded TRs increase the expression of c-fos and c-jun oncogenes, components of the activating protein-1 transcription factor and activate activating protein-1-dependent transcription (50). On the molecular level, to date there are only indirect pieces of evidence that the altered functions of TRs may influence the apoptosis rate in cancer tissues. For example, TRß1 physically interacts with p53 protein, decreasing p53-dependent transcription activation (and, possibly, p53-dependent apoptosis and cell cycle inhibition) (51). In turn, TR-dependent transcription activation also decreases (52). Protein-protein interactions of overexpressed TRs with p53 could excessively intensify these effects.

In light of the results described in this paper (a high frequency of PTCs affected by TR mutations, a high number of mutations within a single TR allele, the functional similarity of mutant TRs to v-erbA), we conclude that TRs, once mutated, could play a role in PTC tumorigenesis. On the other hand, an increased number of thyroid cancers has not been reported in resistance to thyroid hormone patients bearing germline TRB gene mutations. As the direct involvement of mutant TRs in tumorigenesis has not been studied, we can only speculate that yet another factor(s) initializes the process of tumorigenesis, and then mutant TRs exacerbate this process, possibly in part via the molecular mechanisms described above for thyroid tissue, leading to tumor progression and, potentially, to a poorer clinical outcome of the disease.

We also speculate that mutant TRs could be involved in the tumorigenesis of other tissues for the following reasons. TRs are present in all tissues, they directly or indirectly regulate the expression of proteins involved in the proliferation and/or apoptosis control that are present in different tissues (e.g. c-Myc, c-Jun, c-Fos, MDM2, Bax, etc.), they regulate differentiation and maturation (many of their specific direct response genes are known, e.g. the above-described NIS in thyroid tissue, myoD in muscle cells, myelin basic protein in central nervous system, etc.), and the expression of these factors was shown to be altered in cancer tissues. Moreover, LOH of TR genes and their aberrant expression were described in many cancers, and a high frequency of multiple TR mutations affecting receptor function were discovered in other cancer types (25), suggesting that these abnormalities are not specific to PTC. Nevertheless, more information is needed to establish the precise role of mutated TRs in the tumorigenesis of human tissues.

Acknowledgments

We thank Dr. E. Stachlewska-Nasfeter (Department of Brachytherapy, Center of Oncology, Warsaw, Poland) for a kind donation and microscopic evaluation of thyroid tissues. We also thank Dr. J. Pachucki for HEK293 cells.

Footnotes

This work was supported by Polish State Committee for Scientific Research Grant 4P05B04115 (to M.P.-K.).

Abbreviations: CMV, Cytomegalovirus; NFI, nuclear factor I; PBS-T, PBS and 0.1% Tween 20; PTC, papillary thyroid cancer; STM buffer, 0.25 M sucrose, 20 mM Tris-HCl, and 1.1 mM MgCl₂, pH 7.85; SV40, simian virus 40; TRE, thyroid response element.

Received June 6, 2001.

Accepted November 27, 2001.

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