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The Journal of Clinical Endocrinology & Metabolism Vol. 82, No. 12 4229-4233
Copyright © 1997 by The Endocrine Society

Original Studies

Identification of Constitutively Activating Somatic Thyrotropin Receptor Mutations in a Subset of Toxic Multinodular Goiters1

Hans-Peter Holzapfel, Dagmar Führer, Peter Wonerow, Gerhard Weinland, Werner A. Scherbaum and Ralf Paschke

Department of Internal Medicine III, University of Leipzig (H.-P.H., D.F., P.W., W.A.S., R.P.), D-04103 Leipzig; and the Department of Surgery, Israelitisches Krankenhaus Hamburg (G.W.), Hamburg, Germany

Address all correspondence and requests for reprints to: Prof. Dr. R. Paschke, Department of Internal Medicine III, University of Leipzig, Ph.-Rosenthal-Strasse 27, D-04103 Leipzig, Germany.


   Abstract
Top
 Abstract
Introduction
 Subjects and Methods
 Results
 Discussion
 References
 
Constitutively activating mutations in the TSH receptor (TSHR) gene and in the Gs{alpha} gene are frequent molecular causes for solitary toxic nodules of the thyroid. However, the etiology of toxic multinodular goiter is still largely unknown. Therefore, DNA from nodular and quiescent surrounding tissue of six patients with toxic multinodular goiters was screened for mutations in exons 9 and 10 of the TSHR gene and exons 7–10 of the Gs{alpha} gene by direct automated sequencing.

In one patient, two different somatic TSHR mutations were identified in two different toxic nodules (L632I and F631L). In another patient, two different toxic nodules harbored the same TSHR mutation (I630L), whereas only one TSHR mutation (F631L) was identified in one of the two toxic nodules of an additional patient. In the other three patients, no mutations could be found in exons 9 and 10 of the TSHR gene or in exons 7–10 of the Gs{alpha} gene.

Our results demonstrate that not only solitary toxic adenomas but also toxic multinodular goiters can be caused by constitutively activating mutations of the TSHR. In addition to mutations in the TSHR and possibly in Gs{alpha}, there are probably other still unknown mechanisms that cause hot nodules in toxic multinodular goiters.


   Introduction
Top
 Abstract
 Introduction
Subjects and Methods
 Results
 Discussion
 References
 
THE MAIN causes of hyperthyroidism are Graves’ disease and toxic thyroid nodules. In iodine-sufficient areas, toxic nodules are 50 times less frequent than Graves’ disease (1). However, in iodine-deficient areas, autonomous functioning thyroid nodules (AFTNs), classified clinically as solitary toxic nodules (10%) or toxic multinodular goiter (TMG; 48%), account for 58%, whereas Graves’ disease accounts for about 40% of patients with hyperthyroidism (2). AFTNs synthesize and secrete thyroid hormones autonomously, thereby suppressing TSH so that the extranodular tissue becomes functionally quiescent. On the thyroid scintiscan they show increased (hot) radionucleotide uptake compared to that by paranodular thyroid tissue, which has a low uptake due to deprived TSH stimulation.

The coexistence of autonomous and quiescent tissue in the same organ suggests an inherent defect as the cause of AFTNs. This assumption is supported by the persistence of hyperactivity in AFTNs in cell culture and after grafting into nude mice (3). In thyroid epithelial cells the cAMP cascade controls proliferation and differentiated function. Therefore, the clinical observation of hyperthyroidism together with TSH-independent growth of the AFTNs suggest a chronic activation of the cAMP regulatory cascade in AFTNs. Somatic mutations in a gene of the cAMP regulatory cascade leading to constitutive activation of this cascade were first detected in the Gs{alpha} gene in solitary toxic thyroid adenomas (4, 5). Soon thereafter, the first mutations were identified in the third intracellular loop of the TSH receptor (TSHR) in residues homolog to those previously identified in constitutively active mutants of the {alpha}1b-adrenergic receptor (6, 7). A compilation of TSHR mutations identified in solitary toxic adenomas and those found in hereditary or congenital toxic thyroid hyperplasia, shows 16 residues whose mutations to a total of 23 different substitutions confer constitutive activity to the TSHR (6, 8–12, 15–17, 17a) (Fig. 1Go).



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  		Figure 1. Somatic, constitutively activating TSHR mutations identified to date in toxic adenomas and constitutively activating TSHR germline mutations identified in congenital nonautoimmune hyperthyroidism and autosomal dominant nonautoimmune toxic thyroid hyperplasia.

 
However, in contrast to solitary toxic adenomas of the thyroid, the molecular etiology of TMG is still largely unknown. The clinical presentations of solitary toxic nodules and TMGs show many pathophysiological similarities. This suggests a similar reasoning with respect to the role of the cAMP cascade in the etiology of TMGs and that in solitary toxic nodules. To test this hypothesis, the TSHR and the Gs{alpha} gene were sequenced in toxic nodules and quiescent thyroid tissue from TMGs.


   Subjects and Methods
Top
 Abstract
 Introduction
 Subjects and Methods
Results
 Discussion
 References
 
Patients

Thyroid tissue from six patients with TMG was investigated. Diagnosis was based on the clinical finding of thyrotoxicosis with decreased TSH levels, elevated free T3 and/or free T4 values, negative thyroid microsomal and TSHR antibodies, and corresponding results on ultrasound and scintiscan with increased circumscribed technetium uptake by the nodule and suppression of surrounding thyroid tissue. All patients had received antithyroid treatment before surgery. The location and size of the thyroid nodules identified in the six patients are summarized in Table 1Go.


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  		Table 1. Toxic multinodular goiters

 
All experiments were approved by the local ethics committee. Informed consent was obtained from each patient before surgery.

Methods

Toxic nodules were identified by scintiscan, ultrasound, intraoperative inspection, and pathological examination. DNA was extracted from the central part of toxic thyroid nodules and normal periadenomatous thyroid tissue that were collected intraoperatively and immediately stored in liquid nitrogen. Exon 10 of the TSHR was amplified by PCR in 2 fragments of 868 and 875 bp with conditions as follows: an initial denaturation step for 3 min at 94 C, followed by 30 cycles with denaturation for 30 s (95 C), annealing for 30 s (56 C), elongation for 1 min and 30 s (72 C), and a terminal elongation step for 6 min (72 C). 5'-TGG CAC TGA CTC TTT TCT GT-3' and 5'-ACT GTC TTT GCA AGC GAG TT-3' were used as forward primers, and 5'-GTC CAT GGG CAG GCA GAT AC-3' and 5'-GTG TCA TGG GAT TGG AAT GC-3' were used as reverse primers. For amplification of exon 9 of the TSHR the primers were: forward primer, 5'-TCA TCT CCC AAT TAA CCT CAG G-3'; and reverse primer, 5'-GCT TCC AAT TTC CTC TCC AC-3'. Exons 7–10 of the Gs{alpha} were amplified with 5'-TTC TTT TTC TCC CAG CTT CCT-3' as forward primer and 5'-GGT TGG TCT GGT TGT CCT CC-3' as reverse primer. PCR conditions for amplification of exon 9 (TSHR) and exons 7–10 (gsp) were as described above, except for annealing temperatures of 54 C (TSHR) and 52 C (gsp) instead of 56 C. M21-13 and M13 tails were added to all forward and reverse primers, respectively. Sequencing was performed with Thermosequenase (Amersham, Aylesbury, UK) and M21-13 and M13 dye primers (Applied Biosystems, Weiterstadt, Germany). Identification of mutations was also confirmed by sequencing with dye-labeled terminators using the PCR primers as sequencing primers. The sequencing reactions were analyzed with an automatic sequencer (Applied Biosystems 373). All reactions were performed twice. Both strands of the PCR products have been sequenced. All hot nodules summarized in Table 1Go were examined in this way. A positive control with a known Gs{alpha} mutation was included for the Gs{alpha} gene sequencing.

Cloning of the new TSHR mutation

Exon 10 of the TSHR gene was amplified by PCR, using genomic DNA extracted from toxic thyroid nodules (described above) as template. The primers used were as follows: forward primer, 5'-ATC CTT GAG TCC TTG ATG TGT AAT-3'; and reverse primer, 5'-TTA CAA AAC CGT TTG CAT ATA CTC TT-3'. The PCR products were cloned in pUC57 (MBI Fermentas, Vilnius, Lithuania). Resulting recombinant vectors were sequenced with Thermosequenase (Amersham, Aylesbury, UK) and dye-labeled terminators, using the primer 5'-AAG TCC GAT GAG TCC AAC CCG-3', and analyzed with an automatic sequencer (Applied Biosystems 373). Constructs containing the mutant allele were cleaved with CvnI and BstEII (positions 1604–2169). The mutated TSHR constructs were generated by replacing the CvnI-BstEII segment (17) in the wild-type TSHR cloned in pSVl (12a) with the corresponding mutated segment amplified by PCR.

Expression of mutated TSHR constructs

For transient expression in COS-7 cells, the constructs were transfected in 100-mm dishes with 6 µg DNA of wild-type or mutated receptor constructs using the diethylaminoethyl-dextran method (13). Twenty-four hours after transfection, the cells were split and plated in six-well plates. Forty-eight hours after transfection, the cells were used for stimulation and detection of cAMP. Three 30-mm dishes were prepared for each condition.

Measurement of cAMP

Transfected cells (4 x 105/well) were washed with serum-free DMEM without antibiotics after preincubation for 30 min with the same medium containing 1 mmol/L isobutylmethylxanthine. Subsequently, the cells were incubated with or without bovine TSH (100 mU/mL; Sigma Chemical Co., St. Louis, MO) for 60 min in the presence of 1 mmol/L isobutylmethylxanthine. Thereafter, the medium was removed, and 1 mL 0.1 N HCl was added. cAMP was measured in the cell extracts with a commercial kit (Amersham, Braunschweig, Germany) according to the manufacturer’s instructions. The results from a representative experiment are expressed as the mean cAMP values ± SE per 30-mm dish.

Binding assays

Transfected cells (4 x 105/well) were washed once with Hanks’ solution without NaCl containing 280 mmol/L sucrose, 0.2% BSA, and 2.5% low fat milk (12). Thereafter, the cells were incubated in the same medium in the presence of 130,000 cpm [125I]TSH (TRAK Assays, BRAHMS Diagnostica, Berlin, Germany; 25 µCi/µg; 40 U/mg) and the appropriate concentrations of cold TSH at room temperature for 4 h. Before the cells were solubilized with 1 N NaOH, they were washed twice with Hanks’ solution. The bound radioactivity was determined in a {gamma}-counter. TSH or TSHR concentrations were expressed as milliunits per mL. The data were analyzed assuming a 1:1 stoichiometry for TSH binding to its receptor using the fitting module (13a, 14) of SigmaPlot 2.0 for Windows (Jandel Scientific GmbH, Erkrath, Germany).


   Results
Top
 Abstract
 Introduction
 Subjects and Methods
 Results
Discussion
 References
 
In the two toxic nodules located in the left thyroid lobe of patient 1 (Fig. 2Go), two different somatic TSHR mutations were identified, leading to amino acid transitions from phenylalanine to leucine in position 631 and from threonine to isoleucine in position 632 of the TSHR. Both mutations have previously been described in solitary toxic nodules and have shown constitutive activity when expressed in COS-7 cells (15, 16, 17).



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  		Figure 2. Scintiscan of patient 1, showing two hot nodules in the left thyroid lobe in which the Phe631Leu and Thr632Ile mutations were detected.

 
In each of the two hot nodules located in the left thyroid lobe of patient 2, the same new TSHR mutation was found, leading to an amino acid transition from isoleucine to leucine in position 630 of the TSHR (Fig. 3Go). The new mutation was cloned and expressed in COS-7 cells. COS-7 cells expressing the mutated TSHR showed a 5-fold higher basal cAMP accumulation (10.95 ± 0.92 pmol/well) than the wild-type receptor (2.3 ± 0.17 pmol/well), thus demonstrating a constitutive activation of the cAMP cascade (Fig. 4Go). Maximal stimulation with 100 mU/mL TSH showed similar maximal cAMP levels for the mutated TSHR (27.3 ± 0.2 pmol/well) and the wild-type receptor (29 ± 5.8 pmol/well). Binding experiments showed decreased binding capacity for the mutant receptor compared to the wild-type receptor (0.18 ± 0.045 and 0.43 ± 0.09, respectively). The binding capacities exclude overexpression of the mutant receptor as a possible reason for the increased basal cAMP values. The Kd values (2.57 ± 0.5 for the wild-type and 0.8 ± 0.33 for the mutant receptor) showed a higher affinity of the Ile630Leu mutant TSHR for bovine TSH.



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  		Figure 3. TSHR sense sequence for the Ile630Leu mutation detected in two toxic thyroid nodules of patient 2 and the wild-type (WT) TSHR sequence detected in the same patient’s perinodular tissue.

 


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  		Figure 4. Basal and stimulated (100 mU TSH/mL) cAMP values for the Ile630Leu TSHR mutation compared to those for the wild-type (WT) TSHR.

 
In one toxic nodule, located in the left lobe of patient 3, a TSHR mutation was identified with a resulting amino acid transition from phenylalanine to leucine in position 631 of the TSHR. This mutation has previously been shown to be constitutively active when expressed in COS-7 cells (16). No mutation in exons 9 and 10 of the TSHR gene could be found in the other toxic nodule of this patient. The sequence analysis of exons 9 and 10 of the TSHR showed no mutation in the toxic nodules of the patients 4, 5, and 6. In addition, sequencing of exons 7–10 of the Gs{alpha} gene was performed in all nodules in which no TSHR mutation was detected. However, no mutation of the Gs{alpha} protein was found in these exons. Sequencing of TSHR and Gs{alpha} performed for the quiescent thyroid tissue in each thyroid gland showed only the wild-type sequence.


   Discussion
Top
 Abstract
 Introduction
 Subjects and Methods
 Results
 Discussion
References
 
The data presented here provide the first evidence that TSHR mutations are inherent to a major portion of toxic nodules in TMG. Two different and two identical TSHR mutations were detected in the hot nodules of two patients, each with two toxic nodules. One TSHR mutation was found in one of the two nodules in another TMG. Two of the TSHR mutations identified have previously been characterized after transient expression in COS-7 cells and were shown to be constitutively active (15–17, 17a). The new Ile630Leu mutation exhibits constitutive cAMP activity also, as shown by transient expression in COS-7 cells. Sequencing of the perinodular tissue from each case showed only the wild-type TSHR sequence.

There are several possible reasons for the lack of TSHR mutations in three of the six TMGs. Histologically, hypercaptant regions can be classified as either follicular adenomas or adenomatous nodules. The finding that 84% of adenomas but only 33% of adenomatous nodules are monoclonal (18) suggests that this histological heterogeneity correlates with different etiologies. Furthermore, follicular adenomas can be histologically subclassified as micro- or macrofollicular, papillary, trabecular, or atypical (19). In our study there was no correlation between histological phenotype (adenoma or adenomatous nodule) and the presence or absence of a TSHR mutation in the investigated hot nodules. Hot nodules in TMG appear with or without additional nodules that show normal or decreased uptake (cold nodules) on scintiscans. Finally, increased uptake on scintiscan can be circumscribed or patchy, and increased focal uptake can also be detected in up to 50% of euthyroid goiters in iodine-deficient regions (20). Therefore, it is likely that the clinical disorder TMG comprises different and/or overlapping pathophysiological properties that might have different and/or overlapping etiologies, not all of which may be caused by constitutively activating TSHR mutations.

There are several possible explanations as to how and why constitutively activating TSHR mutations might be generated in the evolution of thyroid autonomy. The TSHR seems to be particularly sensitive to mutational events. This is demonstrated by the high number of different TSHR mutations identified to date and the high prevalence of TSHR mutations in solitary toxic nodules (15, 21, 21a). In multinodular goiters there is chronic mutagenic stimulation of the thyroid parenchyma (22) that is likely to provide a sensitive background for mutational events (23, 25). A higher cell number has been reported in TMG compared to normal thyroid tissue (22, 24). This finding would imply increased thyroid epithelial cell proliferation in TMG at least at some time during the evolution of TMG. This would increase the possibility of mutations occurring (25). The increased prevalence of TMG in regions with iodine deficiency (2) suggests that iodine deficiency promotes the mitogenic stimulation of TMG (26). Moreover, additional factors, such as the generation of free radicals associated with the stimulation of the H2O2-thyroperoxidase system (27) or changes in DNA methylation, which have been demonstrated in benign thyroid nodules (28) and are contributing to transcriptional repression and/or point mutations (29), are likely to lead to mutagenic events for which different genes, e.g. the TSHR, seem to be susceptible targets. Although mutations in some expressed genes of the thyroid may appear silent or without easily visible clinical consequences, mutations in the TSHR have been demonstrated as a cause of various thyroid diseases. Further investigations will show whether the TSHR is more susceptible for mutational events than other genes expressed in the thyroid.

In conclusion, constitutively activating mutations of the TSHR seem to be the molecular cause of hyperthyroidism as well as the growth of toxic nodules in a subset of TMGs. Iodine deficiency and the subsequent chronic mitogenic stimulation of the thyroid cell seem to predispose for these mutational events. Due to its susceptibility for mutagenesis, the TSHR appears to be a primary target for mutational events in the thyroid. However, the lack of TSHR mutations in three of six TMGs investigated leaves room for other or additional pathogenic mechanisms involved in the etiology of TMG.


   Acknowledgments
 
We thank U. Scheibler (Surgical Department, Hospital of Dösen, Dösen, Germany) and Dr. P. Lamesch (Surgical Department, University of Leipzig, Leipzig, Germany) for provision of tissue samples. We also thank Dr. H. Kuhn and C. Landmann (Department of Pathology, University of Leipzig) for access to the ABI 373 sequencer for analysis of our sequencing reactions.


   Footnotes
 
1 Presented in part at the 10th International Congress of Endocrinology, San Francisco, CA, 1996. This work was supported by the Deutsche Forschungsgemeinschaft (DFG/Pa 423/3-1) and BMF+F, Interdisciplinary Centre for Clinical Research at the University of Leipzig (01KS 9504, Project B5W). Back

Received November 21, 1997.

Revised July 3, 1997.

Accepted August 15, 1997.


   References
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 Introduction
 Subjects and Methods
 Results
 Discussion
 References
 

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Endocrinology Endocrine Reviews J. Clin. End. & Metab.
Molecular Endocrinology Recent Prog. Horm. Res. All Endocrine Journals