Laboratory of Molecular Biology, National Cancer Institute, 37 Convent Drive MSC 4255,
1 Laboratory of Biochemistry and Genetics, National Institute of Diabetes, Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD 20892-4255, USA,
2 Department of Endocrinology, Medical Research Center, Polish Academy of Sciences and
3 Department of Biochemistry, Medical Center of Postgraduate Education, Warsaw, Poland
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Abstract |
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Abbreviations: EMSA, electrophoresis mobility shift assay; HCC, hepatocellular carcinoma; LOH, loss of heterozygosity; NFTs, non-functioning tumors; RCCC, renal clear cell carcinoma; RTH, thyroid hormone resistance syndrome; RXRs, retinoic X receptors; T3, thyroid hormone; TRs, thyroid hormone nuclear receptors; TRE, thyroid hormone response element; VHL, von HippelLindau tumor suppressor gene; w-TR, wild-type TR.
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Introduction |
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Thyroid hormones regulate growth, development and differentiation. Their action is mediated by the thyroid hormone nuclear receptors (TRs), which are derived from two genes, TR and TRß, located on chromosomes 17 and 3, respectively (10). Each gene gives rise to two receptor isoforms,
1,
2, ß1 and ß2, by alternative splicing of the primary transcripts. The gene regulating activity of TRs is mediated by binding to specific DNA sequences, known as thyroid hormone response elements (TREs), located on the promoter regions of thyroid hormone target genes. Recent studies have indicated that the gene regulatory functions of TR are modulated by formation of different heterodimers with other members of the receptor superfamily, notably the retinoic X receptors (RXRs) (10,11), by differential interaction with different types of TREs (10) and by interaction with various co-activators, co-repressors and other cellular proteins (10,12).
Increasing evidence has suggested that aberrant expression and/or mutations in TR genes could be associated with carcinogenesis. A reduction in the expression of mRNA for TRß1 and TRß2 was implicated in inappropriate expression of the glycoprotein hormone -subunit gene in non-functioning tumors (NFTs) of the anterior pituitary and was proposed to contribute to uncontrolled tumor growth (13,14). Reduced expression of TRß1 was also found in poorly differentiated fibroblast-like osteosarcoma (15). However, in poorly differentiated hepatocarcinomas, overexpression of TRß1 was correlated with enhanced thyroid hormone (T3)-induced proliferation (16,17). These results suggest that aberrant expression of TRs may be associated with different types of tumors and/or different states of differentiation. Furthermore, LOH of the chromosomal regions where TR genes are located (3p21p25 for the TRß gene and 17q21 for the TR
gene) was a frequent event in tumors (1821). For example, LOH of the TRß gene was observed in most of the small cell lung cancers examined (20,22), in 60% of posterior uveal melanomas (23), 30% of breast (24) and 64% of non-familial renal cell carcinomas (25). Consistent with these observations, microdeletion of both TR
alleles was found in 20% of gastrointestinal tumors (26). LOH of the TR
gene was also observed in 79% of breast (27) and prostate cancers (28). In addition to LOH in regions in which TR
and TRß genes are located, high frequencies of mutant TRs (65% of TR
1 and 76% of TRß1 in 17 tumors) with impaired function were identified in human hepatocellular carcinoma (17). TR
1 mutants (13% of 23 tumors) were also found in NFTs of the anterior pituitary (14).
We had previously found aberrant expression of TR and TRß mRNAs in RCCC (29). Expression of both TR
1 and TR
2 mRNAs was reduced, while TRß1 mRNA was overexpressed in 30% and significantly reduced in 70% of tumors examined. We hypothesized that these aberrant expression patterns may reflect mutations in TR genes leading to an alteration in expression. In the present study we have cloned TR cDNAs from tumors obtained from RCCC patients. We found multiple mutations in both the TR
and TRß genes, resulting in impairment of T3 and DNA binding and loss of transcriptional activity. The functional impairment in these mutants may contribute to the carcinogenesis of RCCC.
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Materials and methods |
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Tissues
Tissues were kindly provided by Dr A.Tanski of the Department of Urology, Vouvodoship Hospital (Ostroleka, Poland) with the permission of the Ethical Committee on Human Studies, Medical Center of Postgraduate Education, during unilateral nephrectomies following the diagnosis of kidney cancer. Fragments of the tumors, fragments of the opposite poles of the same kidney (not infiltrated with cancer) and fragments of the kidneys with no cancer were excised, immediately frozen on dry ice and stored at 75°C until needed. In total, 22 tumor tissues, 20 corresponding controls and seven non-cancerous kidney tissues were collected. Upon histological evaluation, the renal clear cell cancer diagnosis and tumor borders were established. Tumors were divided into the three groups depending on the grade of differentiation: G1, well differentiated (four tumors); G2, moderately differentiated (10 tumors); G3, undifferentiated cancers (eight tumors). Additionally, they were divided according to the TNM (tumor, nodules and metastases) classification of malignant tumors (31).
RNA isolation
RNA was isolated according to Chomczynski (32). Up to 100 mg deep frozen kidney tissue was manually homogenized in a glassteflon homogenizer directly in 1 ml of TRI reagent (Molecular Research Center, Cincinnati, OH). Samples were incubated at room temperature for 510 min, supplemented with 0.2 ml of chloroform, mixed by shaking for 15 s, incubated at room temperature for 10 min and then centrifuged at 12 000 g for 15 min at 4°C. The upper, aqueous phase was transferred to another Eppendorf tube, mixed with 0.5 ml of isopropyl alcohol, incubated at room temperature for 15 min and then centrifuged at 12 000 g for 15 min at 4°C. The RNA pellet was washed with 75% ethanol, air dried and resuspended in 4050 µl of DEPC-treated water. Purity of the RNA was evaluated by formaldehydeagarose electrophoresis and the concentration determined by spectrophotometric measurements.
Cloning of TR cDNAs from kidney cancer tissues
To clone TR1 and TRß1 cDNAs in kidney cancer tissues as well as in normal tissues, RTPCR was performed using a Superscript One-Step RT-PCR System (Life Technologies, Grand Island, NY) or Ready To Go RTPCR beads (Amersham Pharmacia Biotech). Occasionally the reactions were supplemented with magnesium to a final concentration of up to 3 mM. To clone TR cDNAs, the following primers were used: for TR
1, forward primer 5'-GGATGGAATTGTGAATG-3' or 5'-ATGGAACAGAAGCCAAGCAA-3' and reverse primer 5'-GGCCGCCTGAGGCTTTA-3'; for TRß1, forward primer 5'-GATCCAGAATGATTACTAACC-3' and reverse primer 5'-GGAATTATAGGAAGGAA TCC-3' and then internal primers, forward 5'-CTATAACCCCCAACAGTATG-3' or 5'-ATGACAGAAAATGGCCTTAC-3' and reverse, 5'-CTAATCCTCGAACACTTCCA-3'. Total RNA (0.51 µg) was used as a template in each RTPCR reaction. Prior to each RTPCR, RNA was denatured at 80°C for 5 min, cooled to room temperature and then supplemented with enzyme mix. Reverse transcription was performed at 48 or 4243°C for 30 min. Reverse transcriptase was inactivated by incubation of the samples at 94°C for 23 min. cDNA amplification was performed using the following conditions: 40 cycles of 94°C for 4050 s, 52°C for 11.5 min and 72°C for 2.5 min, followed by 10 min incubation at 72°C. In addition, since the amount of TRß1 was very low after RTPCR, a second PCR was performed with the above described internal primers. RTPCR products were electrophoresed on 1% agarose gels, then the gel fragment at the expected TRß1 size was excised, dissolved in 100 µl of dH2O and 5 µl of this solution was used as template in a second PCR: 3 min at 94°C, 25 cycles of 94°C for 4050 s and 52°C for 1 min, then 72°C for 2 min, followed by 10 min incubation at 72°C.
TRß1 or TR1 cDNAs derived from RTPCR were electrophoresed on 1% agarose gels, excised and extracted from the gel with a QiaQuick Gel Extraction Kit (Qiagen, Valencia, CA), and then ligated into vector pGEM-T containing T overhangs (Promega, Madison, WI). JM 109 bacteria were transformed with the ligation mix and bluewhite selection was performed. Mini-preps were carried out using the Wizard Plus SV Minipreps DNA Purification System (Promega, Madison, WI) or Plasmid Miniprep Plus kits (A&A Biotechnology, Gdansk, Poland). Restriction analyses were carried out to confirm the correct cloning (TR
1 with NcoI and TRß1 with BstXI) in the pGEM-T vector. In the preliminary studies we have validated the above cloning procedure by isolating wild-type TRs from nomal kidneys and confirmed the sequences of wild-type TRs by sequencing.
Identification of mutation sites by sequencing of TRs isolated from kidney cancer tissues
To identify mutations in TRs isolated from tissues, the TR cDNAs cloned in the pGEM-T vector were sequenced by automatic sequencing using a BigDye Terminator Cycle Sequencing Kit (Perkin Elmer) using T7 and SP6 as primers. Mutation was confirmed by repeated sequencing of the same clone as well as other clones (up to four) originating from the same tumor. Sequence analysis was examined with Lasergene software (DNAstar, Madison, WI). As a control, wild-type TRß1 was cloned from non-cancerous kidneys and sequenced to validate the cloning and to confirm the accuracy of sequencing reactions.
Cloning of TRß1 and TR1 mutants into pBluescript KS()
For efficient in vitro transcription/translation of mutant proteins, three TR1 mutants (23TR
1, 2TR
1 and 6TR
1) and one TRß1 mutant (18TRß1) were recloned into the pBluscript KS() expression vector, which utilizes T7 as the promoter. The TR cDNAs were cut from the pGEM-T vector with the restriction enzymes SpeI and ApaI and cloned into the same sites in the pBluscript KS() expression vector, which was confirmed by restiction analyses using BstXI or NcoI for TR
1 and BstXI or PstI with SpeI for TRß1.
Cloning of the TRß1 and TR1 mutants into a eukaryotic expression vector
The same TR1 and TRß1 mutants described above, positioned with their 5'-ends next to the SP6 promoter within the pGEM-T vector, were cut from this vector with NotI and ApaI and subcloned into vector pcDNA 3.1(+) (Invitrogen, Carlsbad, CA), previously prepared with the same enzymes. To confirm correct cloning of the inserts, restriction analysis was performed with NcoI (TR
1 clones) or BstXI (TRß1 clones). The remaining six mutated TRß1 mutants, positioned with their 5'-ends next to the T7 promoter within the pGEM-T vector, were restricted with NcoI, treated with Klenow fragment (to blunt ends), phenol/chloroform and gel purified, then restricted with NotI and cloned into pcDNA 3.1(+), previously prepared with EcoRV and NotI. Restriction analysis was performed with BstXI.
These mammalian expression plasmids were purified using the Qiagen Maxi Kit and resequenced to confirm the mutation sites determined above. The sequencing was carried out using an Applied Biosystems model 377 automatic DNA sequencer according to the manufacturer's instructions (Applied Biosystems, Foster City, CA).
Preparation of nuclear extracts from kidney cancer tissues
The buffers used for isolation were as described by Kane (33). All buffers were supplemented with pepstatin A (final concentration 1 µg/ml), leupeptin (1 µg/ml), aprotinin (2 µg/ml) and PMSF (0.5 mM). Up to 100 mg of the tissue was manually homogenized in a glassteflon homogenizer in 1 ml of ice-cold STM buffer (0.25 M sucrose, 20 mM TrisHCl, pH 7.85, 1.1 mM MgCl2) and centrifuged at 1000 g for 10 min at 4°C. The pellet was washed twice in 1 ml of STM buffer with 0.5% Triton X-100, followed by centrifugation under the conditions described above. The final pellet was resuspended in 0.1 ml KSTM + 20% glycerol buffer (0.25 M sucrose, 20 mM TrisHCl, 1.1 mM MgCl2, 0.4 M KCl, 20% glycerol, 5 mM DTT), sonicated and then incubated on ice for 30 min with vortexing every 5 min to extract the soluble nuclear proteins. After solubilization, the suspension was centrifuged at 12 000 g for 15 min at 4°C. The amount of protein in the supernatant was quantified by spectrophotometry with Bio-Rad Protein Assay buffer at 595 nm and stored in 10 µl aliquots at 75°C.
Electrophoresis mobility shift assay (EMSA)
TR proteins were synthesized by in vitro transcription/translation using the TNT coupled reticulocyte kit according to the manufacturer's instructions (Promega). 32P-labeled TRE-Lys was prepared as previously described (30). The in vitro translated TR proteins were quantified by measuring the intensity of the 35S-labeled protein bands after SDSPAGE. For EMSA, identical amounts of TRs were incubated with the 32P-labeled TRE in the presence or absence of RXRß. After electrophoresis, TR homodimers and heterodimers were visualized by autoradiography.
To further evaluate whether the DNA-binding activity of TRs was altered in RCCC, gel mobility shift assays were performed with 7.510 µg nuclear extract isolated from tumors and their respective controls. The probe was a double-stranded DNA containing TRE-DR4 (5'-GATCGCAGGTCATTTCAGGACAGCGATC-3'). Mutated TRE served as a non-specific competitor (5'-GGCAAATCATTTCAAGACAG-3'). Nuclear extracts were incubated at room temperature for 20 min in binding buffer containing 20 mM TrisHCl, pH 7.5, 50 mM KCl, 2 mM DTT, 0.1% Triton X-100, 6% glycerol, in the presence of 250 ng dI·dC, 1 ng probe and a 10 times excess of specific or non-specific competitor. For supershift experiments, 1 µg mouse monoclonal anti-TR (ß1 and 1) antibody C4 (34) was added to the binding reaction.
The mixture was first incubated on ice for 30 min, followed by an additional incubation at room temperature for 20 min. The binding reactions were loaded onto a 4% native gel and electrophoresed at 150 V for 2 h at room temperature. The gel was dried and autoradiographed.
Binding of [3'-125I]T3 to TRs
The in vitro translated TR proteins were incubated with 0.2 nM [3'-125I]T3 in the presence of increasing concentrations of unlabeled T3. The TR-bound [3'-125I]T3 was separated from free [3'-125I]T3 as described by Zhu et al. (30). The binding data were analyzed using equation 1 (below), based on direct competition between [3'-125I]T3 and unlabeled T3 for a single site on the receptor. The concentration of radioactive complex is given by:
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Determination of the transactivation activity of TRs
To determine the T3-dependent transactivatioin activity, CV1 cells (6-well plates with 1x105 cells/well) were transfected with mammalian expression plasmids for mutant (mutant TRß1, 0.21.6 µg; mutant TR1, 0.40.6 µg) and/or wild-type TR (TRß1, 0.050.2 µg; TR
1, 0.1250.5 µg) and 0.8 µg TRE-containing luciferase reporter gene (pTK-Pal-Luc or pTK-Lys-Luc) by the Lipofectamine method according to the manufacturer's instructions. After 16 h the medium was changed to a medium containing 10% thyroid hormone-depleted fetal bovine serum with or without 100 nM T3. After 24 h the cells were harvested and lysed and 100 µl was assayed for luciferase activity according to the manufacturer's instructions (BD PharMingen, San Diego, CA). The transfection efficiency was normalized to the protein concentration of the lysates.
Western blotting
Cell lysates (75 µg) from transient transfection experiments, as described above, were loaded onto a 10% SDSPAGE gel. After electrophoresis, proteins were transferred to a PVDF membrane. The membrane was gently shaken in 10% non-fat milk in Tris-buffered saline (25 mM Tris, pH 7.4, 150 mM NaCl) for 1 h and subsquently washed three times with Tris-buffered saline. The membrane was incubated with mouse monoclonal antibody J51 (2 µg/ml) (36) or C4 (2 µg/ml) (34) overnight at 4°C. After washing with washing buffer (0.1% Tween 20, 25 mM Tris, pH 7.4, 150 mM NaCl), the membrane was incubated with rabbit anti-mouse Ig conjugated to horseradish peroxidase (1:2000 dilution). TR protein bands were visualized by chemiluminescence using the ECL kit (Amersham Pharmacia Biotech).
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Results |
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Examination of the mutations shown in Figure 1 indicates that single (8TRß1 and 25TRß1) and multiple mutations (3TRß1, 18TRß1, 15TRß1, 6TRß1, 32TRß1 and all TR
1 mutants) were detected. In addition to the S380F mutation, 32TRß1 had a 26 amino acid deletion at the N-terminus. The majority of mutations were clustered in the hormone-binding domain (domains D + E, see Figure 1
). This mutation pattern is similar to that observed for TR mutants isolated from human hepatocellular carcinoma (HCC) in that single and multiple mutations were observed (17). This is in contrast to that seen for mutations identified for patients with the genetic disease thyroid hormone resistance syndrome (RTH) (35) in which only a single mutation for each mutant was found in the TRß gene. Analysis of the mutation patterns of RTH patients showed that the mutations are clustered in three regions in the T3-binding domain (36). It is of interest to point out that F451I in 3TRß1, F451S in 8TRß1, L456S in 6TRß1 and M388I in 2TR
1 and 6TR
1 occurred in the first `hot-spot'. The Y321H mutation identified in 25TRß1 and K288E in 23TR
1 were in the second hot-spot and H184Q, S183N and R228H in 23TR
1 and A225T in 6TR
1 were in the third hot-spot identified in RTH patients (36). However, novel mutations were also observed in the DNA-binding domain for 3TRß1 (S99R), 15TRß1 (K155E), 2TR
1 (I116N) and 6TR
1 (I116N) (Table I
).
T3-binding activity of TR mutants is impaired
To understand the functional consequences of mutations in the TRs isolated from RCCC, we first evaluated hormone-binding activity of TR mutants. Mutant TR proteins were prepared by in vitro transcription/translation. Except for 32TRß1, which had an apparent molecular mass of 52 kDa due to deletion of 26 amino acids at the N-terminus (Figure 2A, lane 8), all other TRß1 mutants had sizes similar to w-TRß1 (Figure 2A
, lane 1 versus lanes 27). The in vitro translated wild-type and mutant TRß1 were full-length receptors with an apparent molecular mass of 55 kDa together with a smaller protein, most likely due to initiation from a downstream ATG (36,37). The full-length in vitro translated TRß1 mutants were recognized by monoclonal antibody J51, whose epitope is located in the second half of the A/B domain of TRß1 (36). The 52 kDa 32TRß1 was recognized by monoclonal anti-TR antibody C4, whose epitope is the C-terminal 457EVFED461 of TRß1 and TR
1 (data not shown; 34). Taken together, the identity of the cloned TRß1 mutants was confirmed.
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To determine T3-binding activity, equal amounts of in vitro translated wild-type and mutant TRs were used for binding to [125I]T3. Figure 3 shows the competitive binding displacement curves for wild-type and mutant TRs. Analyses of the binding data shown in Figure 3A
indicate that w-TRß1 bound to T3 with a Kd of 0.78 nM (Table II
). The mutations in 15TRß1 (K155E and K411E) did not affect its T3-binding activity as 15TRß1 retained the hormone-binding activity of w-TRß1. The mutations in the hormone-binding domains of 25TRß1 (Y321H), 6TRß1 (E299K, H412R and L456S) and 32TRß1 (S380F) reduced T3-binding activity by 35, 60 and 46%, respectively (Table II
). The mutations in the hormone-binding domains of 3TRß1 (W219L and F451I), 8TRß1 (F451S) and 18TRß1 (Q252R, A387P and F417L) led to a >99% loss of T3-binding activity. A similar analysis of the data shown in Figure 3B
indicates that the mutations in 23TR
1 (S183N, H184Q, R228H and K288E), 2TR
1 (M388I) and 6TR
1 (A225T and M388I) resulted in virtually complete loss of T3-binding activity (Table II
).
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To determine the DNA-binding characteristics of endogenous TRs in cancer tissues, nuclear extracts of RCCC tissues as well as extracts prepared from the healthy opposite kidney poles were evaluated by EMSA assay (Figure 5A). The DNA-binding patterns were analyzed in 20 cancercontrol pairs. Binding of the TRE to TRs present in nuclear extracts of RCCC in which mutations were found was much weaker than that in the healthy controls. One representative example (a tumor from patient 18) in which no binding was detected is shown in lane 2 (Figure 5A
, lanes 2 versus 5). No significant changes in DNA binding were found in the tumor nuclear extracts from patient 32 (see Figure 5A
, lane 8 versus lane 11). The DNA-bound bands shown in lanes 5, 8 and 11 were specific because the intensities of the bands were competitively reduced in the presence of a 10-fold excess of unlabeled TRE (lanes 5, 8 and 11 versus 7, 10 and 13, respectively). No competition was seen in the presence of non-specific competitors (lanes 5, 8 and 11 versus 6, 9 and 12, respectively).
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The T3-dependent transactivation activity of TR mutants is impaired
To further assess the functional consequences of mutations in TRs isolated from RCCC, we determined the transactivation activity of TR mutants by transient transfection assays. Mammalian expression plasmids for the mutants were co-transfected with TRE-containing reporter genes into CV-1 cells. Figure 6A compares the transactivation activity of TRß1 mutants using a reporter containing TRE-Lys. In the absence of T3 all mutants were more potent than w-TRß1 in repression of basal transactivation activity (bar 3 versus bars 5, 7, 9, 11, 13, 15 and 17). Except for 32TRß1, the T3-dependent transactivation activities of all other mutants were reduced, ranging from 50% reduction as seen for 25TRß1 (bar 12 versus bar 4) to a total loss of transactivation activity for 8TRß1 (bar 8). A similar reduction in transactivation activity was detected using the reporter TRE-Pal (data not shown).
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Similar to the observations for TRß1 mutants, all three TR1 mutants were more potent repressors than w-TR
1 in the absence of T3 (Figure 6B
, bar 3 versus bars 5, 7 and 9). Furthermore, the T3-dependent transactivation activities of all three mutants were nearly completely lost (bars 6, 8 and 10). Figure 6D
indicates that the loss of transactivation activity of TR
1 mutants was not due to lower expression of TR
1 mutant proteins because the levels of expression of the TR
1 mutant proteins were comparable to that of w-TR
1. Thus, the loss of transactivation activity was the consequence of loss of T3-binding and/or DNA-binding activity (Table II
).
Dominant negative action of TR mutants
We further evaluated the dominant negative action of TR mutants by examining their inhibitory effect on the transactivation activity of w-TRs. CV-1 cells were transfected with the w-TR expression plasmid together with a 5- or 10-fold excess of mutant expression plasmid. The transactivation activities of w-TR in the absence or presence of mutant TR were compared and the results are shown in Table III. 8TRß1 was a strong dominant negative mutant in that 49 and 84% of the transactivation activity of w-TRß1 was inhibited at mutant:w-TRß1 plasmid ratios of 5:1 and 10:1, respectively. 3TRß1 exhibited a dominant negative effect at a T3 concentration of 10 nM. However, at a 10-fold higher T3 concentration the dominant negative effect of 3TRß1 was abrogated. This is consistent with the T3 binding affinities of these two mutants (see Table II
). All other TRß1 mutants lacked dominant negative activity (Table III
). The three TR
1 mutants, 23TR
1, 2TR
1 and 6TR
1, exhibited strong dominant negative activity in that they inhibited 6379 and 8186% of the transactivation activity of w-TR
1 at mutant:w-TR
1 plasmid ratios of 5:1 and 10:1, respectively (Table III
).
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Discussion |
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We therefore cloned TR cDNAs from RCCC and evaluated their functions. Sequence analyses of TR cDNAs isolated from RCCC show that the TRß and TR genes were mutated in seven of 22 and three of 22 tumors, respectively. These somatic mutations led to loss of the hormone-binding, DNA-binding and transcriptional activities. The extent of functional impairment depends on the sites and number of mutations in the TRs. The molecular basis of the functional impairment of these mutant TRs, however, is difficult to quantify. Most contain several mutated residues; the contributions of these changes may not be additive (42). Furthermore, we have shown that T3-dependent transcriptional activation of TRs is modulated by interactions between all domains in the protein (30). Consequently, it is difficult to predict the results of any of these changes. Interestingly, some of the mutations identified in RCCC occurred in the three `hot-spots' in the hormone-binding domain of TRß1 identified from patients with RTH (36). Thus, 8TRß1 has a single mutation (F451S) in hot-spot 1 and has greatly reduced affinity for T3 (Table II
). However, 25TRß1, which has a single mutation (Y321H) in hot-spot 2, exhibits only a minor decrease in its affinity for T3, but had only 50% of the wild-type binding to the TRE. 15TRß1 has two sequence changes, one in the DNA-binding domain (K155E) and the other in the putative dimerization interface (K411E) (43). Consistent with these changes, this mutant retains affinity for T3 but shows no binding to DNA. Two of the mutations in 6TRß1 occur at the dimer interface (H412R) and near hot-spot 1 (L456S), respectively, resulting in a significant loss of binding to DNA and a minor reduction in T3 binding. A significant reduction in T3 binding is seen for 18TRß1, which includes sequence changes in hot-spot 3 (Q252R) and the dimerization domain (F417L). Alterations in 3TRß1 were found in hot-spot 1 (F451I), which is consistent with a large decrease in affinity for T3.
Genetic changes in the TR1 mutants are equally complex. 2TR
1 and 6TR
1 show similar behaviour, presumably due to the same structural changes, a large reduction in affinity for the hormone and loss of homodimer binding to TRE-Lys (I116N is in the last helix of the DNA-binding domain). The very low affinity for hormone shown by 23TR
1 is readily explained by the four mutations found in its ligand-binding domain (three in hot-spot 3 and one in hot-spot 2). However, the origin of its reduced capacity to bind to DNA is far from clear and awaits elucidation.
TR1 and TRß1 with multiple mutations were also found in HCC (17). All mutated TR
1 identified in HCC with known mutation sites (eight of 12 HCC evaluated) were found to have between two and four mutations. For TRß1 mutants, 50% from HCC were found to have between two and four mutations. This is in contrast to RTH patients, in which no multiple mutation of the TRß gene was found and, moreover, no mutation was detected in the TR
gene. The role of multiple mutations of TRs in carcinogenesis is yet to be elucidated.
Comparison of the mutation sites in TR1 and TRß1 mutants between RCCC and HCC (17) shows that there was only one common mutation site in TR
1, at A225, and no common mutation site was detected in TRß1 mutants between these two cancers. In RCCC, A225 of TR
1 was mutated to T, whereas in HCC A225 was mutated to G. However, in spite of different mutations of TRs in these two cancers, functional impairment of mutant TRs derived from these two cancers was similar in that mutation led to loss of the hormone-binding, DNA-binding and transcriptional activity. Moreover, TR mutants frequently exhibited dominant negative activity (17,44,45). These findings suggest that impairment of TR functions could play a critical role in the tumorigenesis of these two cancers.
At present, however, the precise roles of mutant TRs in carcinogenesis are not clear. Based on the functional impairment of TRß1 and TR1 mutants demonstrated in the present study, it is reasonable to postulate that the transcriptional regulation of TR-mediated genes involved in cell differentiation, proliferation (4648) and apoptosis (4951) is abnormally affected in RCCC expressing these mutants. In addition, functional impairment of TRß1 and TR
1 mutants might also affect the proteinprotein interactions in the network of cellular proto-oncogenes and tumor suppressors. TRs have recently been found to cross-talk with other signalling pathways. For example, TRß1 was shown to physically associate with the tumor suppressor protein p53 and repress p53-mediated transcriptional activity. Conversely, p53 reduces the ability of TRß1 to bind to DNA and to activate transcription (52,53). TRs also stimulate expression of the c-fos and c-jun proto-oncogenes, increase expression of the c-Fos and c-Jun proteins and activate AP1 transcriptional activity in a T3-independent pathway. In turn, both c-Fos and c-Jun inhibit T3-dependent transcription activation (5456). TRs have been shown to directly activate expression of the mdm2 oncogene, which subsequently induces rapid degradation of p53 (57,58) and inhibits retinoblastoma tumor suppressor. Thus, functional impairment due to mutations and the aberrant expression of TRs in RCCC could result in the disarray of the normal regulatory control of cell proliferation, differentiation and apoptosis. Thus, dedifferentiated cells could again become sensitive to proliferation signals, while cells with damaged DNA are possibly not eliminated by means of apoptosis. The present study demonstrates that the percentage of mutated TRß1 was lowest in well differentiated (G1) and highest in poorly differentiated and fast growing (G3) tumors (see Table I
). This finding is consistent with the notion that regulatory control of these processes is severely affected in poorly differentiated tumors. The functional consequences of mutations may be further accentuated by the dominant negative action of mutants. Indeed, our data clearly indicate that 3TRß1, 8TRß1, 23TR
1, 2TR
1 and 6TR
1 exhibit potent dominant negative activity. In addition to the loss of and/or interference with the normal functions of TRs, TR mutants may act via a gain-of-function mechanism. This mode of action of mutant genes is not unprecedented, as it has been clearly documented in p53 mutants (59). The gain-of-function of TR mutants may provide a growth advantage early in the progression of neoplastic cells, may affect genes which prevent differentiation and/or apoptosis of cells and may promote genomic instability. Any of these actions may contribute to the development of RCCC. These possibilities will await validation in future studies.
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Notes |
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* The first two authors contributed equally to this work.
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Acknowledgments |
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References |
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