Since the discovery of TSHR-Ab as the causative agent of GD, there have been several studies on the significance of the measurement of these Ab during the course of the disease and during antithyroid drug treatment.13,14 In general, there are 2 methods of detecting Ab to the TSHR: 1) competitive-binding assays and 2) cell-based bioassays. The conventional binding assays only yield information about the presence or absence of Ab and their concentrations. Bioassays yield more information. These assays determine the function of the antibody, either stimulatory or inhibitory (blocking). Therefore, antibody binding assays only report the presence or absence of TSHR-Ab and their concentrations but do not indicate their functional activity. Bioassays, in contrast, indicate whether TSHR-Ab have stimulatory or blocking activity.
In this review, we will provide a historical summary of the various methods used to detect the functionality of TSHR-Ab and describe some of the recent advances in bioassay technology. We will also discuss the clinical relevance of both stimulatory and blocking activity of TSHR-Ab. Recent iterations of bioassays are offered by clinical reference laboratories; however, in the near future, they will become more widely available to clinicians and more standardized. Finally, continued improvements in these sensitive assays will help to facilitate their routine performance by clinical laboratories.
There is a variety of terms used to describe the different types of TSHR-Ab.15 Thyroid stimulating hormone receptor antibodies refers to any type of Ab specific to the TSHR, but it is commonly used in reference to Ab detected in an immunoassay. Most immunoassays today use a competitive-binding assay and measure what are referred to as TSHR-binding inhibitory immunoglobulins (TBII). In contrast, bioassays measure either TSI, also referred to as TSHR stimulatory antibodies, or TSHR-blocking immunoglobulins also called TSHR-blocking antibodies. Alternative terminologies for blocking antibodies are TSHR-stimulating blocking antibodies and TSHR-blocking antibodies. To avoid possible confusion, in this review, we will use TSHR-Ab as a general term to refer to anti-TSHR-Ab regardless of how they are measured. We will use TBII to refer to the antibodies measured via binding assays, whereas antibodies measured via bioassays will be referred to as TSI for stimulatory.
DETECTION OF TSHR-Ab
Competitive-Binding Assays for TSHR-Ab.
Patient sera are tested for the ability of Ab to compete with either TSH or TSHR monoclonal antibodies (MAb) to bind to intact TSHR or a fragment of the TSHR.16 The first receptor assay for TSHR-Ab was described in 1974.17 Solubilized human thyroid membrane preparations and 125I-labeled bovine TSH (bTSH) were incubated with test material. After incubation, the reaction mixture was centrifuged, and bound 125I-labeled bTSH was counted. The procedure required the precipitation of patients immunoglobulin G (IgG) prior to running the assay. An upgraded version of this receptor assay for TSHR-Ab was introduced in 1982.18 The procedure used detergent-solubilized porcine TSH receptors, 125I-labeled TSH and polyethylene glycol-precipitated IgG from patient sera.
The production of MAb in the late 1990s led to development of second generation TBII assays. Porcine TSHR were immobilized with the mouse MAb 4E31,19 and recombinant human TSHR were immobilized with the MAb BA8.20 Further, these solid-phase assays used either an Enzyme-Linked Immunosorbent Assay format or coated polystyrene tubes and MAb to immobilize the TSHR on the wells of a multiwell plate.20,21 The TBII assays were no longer radioactive and used either chemiluminescent or peroxidase readout.19,20 In second generation assays, TBII were shown to be detectable in approximately 95% of untreated patients with GD.22
The third generation TBII assays are widely distributed and commercially available and measure the ability of Ab in patient serum to inhibit the binding of labeled F(ab)2 human TSHR-stimulating MAb, M22, to TSHR.23 In a described automated version of this assay, a rubidium-labeled F(ab)2 fragment of M22 is used with porcine TSHR bound to plastic tubes, and competitive binding is measured.24 The third generation TBII assays enable the detection of TSHR-Ab of approximately 95% to 98% of untreated patients with GD.25,26
Binding Versus Functional Bioassays for TSHR-Ab.
Today, many variations of TSHR-Ab assays are offered; however, all assays are either immunoassays that measure binding (TBII) only or bioassays that differentiate the Ab functionality. Both, TBII assays and functional bioassays have improved greatly during the last decade. Lack of assay standardization has been a problem in the past. This situation has improved with the availability of international standard material and the use of anti-TSHR MAb in place of TSH as calibration material. For TBII assays, this development has resulted in a close correlation among assays from different manufacturers,25 and assay turnaround times are now under 2 hours. The most recent iterations of TBII and functional assays show clinical sensitivity and specificity of >90% for the diagnosis of GD versus other causes of hyperthyroidism.24,25 In children, these assays have historically been shown to be less accurate for detection of GD, with 60% sensitivity,27 but modern assays now show comparable sensitivity and specificity to what is observed in adults.28,29
As to differences between TBII assays and functional bioassays, comparisons of their dose-response curves after application of serial dilutions of international TSHR-Ab standard material suggests that functional bioassays might be more sensitive for diagnosing subtle disease, at the eventual expense of accuracy at very high TSHR-Ab concentrations.30 In contrast to the TBII assays, the biological activity of specific IgG is directly assessed with the bioassay on a fully functional TSHR holoreceptor expressed on intact live cells, a platform that is easily adaptable and tailored to detect Ab of specific function. As shown in Table, further differences, advantages, and disadvantages of TBII versus functional assays are compared and discussed.
CELL-BASED BIOASSAYS FOR TSI
Chinese Hamster Ovary Cells.
The cloning and sequencing of the TSHR in 198939 enabled the development of stably transfected eukaryotic cell lines expressing the human recombinant TSHR.40 The highest TSHR numbers per cell were found in the cell lines Jason Perret (JP) 14, JP26, and JP28. The original JP series of clones provided a basis for the development of novel bioassay methods for TSI.40 The standard procedure for the measurement of cyclic adenosine 3′, 5′-monophosphate (cAMP) accumulation in Chinese hamster ovary (CHO) cells transfected with the human recombinant TSHR required serum IgG purification with polyethylene glycol and suspension in Hanks balanced salt solution or in hypotonic buffer.
To make bioassays more suitable in a clinical routine laboratory, cryopreserved stocks of CHO cells subclone K1 and CHO JP09 cells transfected with the recombinant human TSHR were used to measure the stimulatory activity in patients with GD.41,42 The use of frozen-thawed stocks of cells influenced their growth rate and the production of cAMP.42 The assay was performed with the CHO cell line JP09 expressing the recombinant TSHR in a radioimmunoassay.41 The major advantage of this system was the use of unfractionated serum which avoided the troublesome IgG purification.41 Another assay method used also the CHO cell lines, clone JP02, and clone JP09, and additionally the CHO cell line JP26 both expressing the recombinant human TSHR.43 Further in this work, the binding capacity of bTSH was compared for the 2 cell lines by substitution of 125I-labeled bTSH when using rising concentrations of unlabeled bTSH. As a result, the bTSH binding capacity was much higher in the JP09 cell line compared with the JP26 cell line. The CHO JP09 cells displayed a higher stimulation index when exposed to bTSH than JP26 cells due to the lower number of expressed human TSHR in the JP26 cell line. The JP09 cell line expressed 90,000 versus 2,000 human TSHR per cell only for the JP26 cell line. Human thyroid epithelial cells express between 1,000 and 10,000 TSHR per cell, but most of the experiments used the JP09 cell line. The cell numbers also varied between the different investigators.
The first reporter gene-based bioassay for the detection of TSI used the CHO cell line C6-13 stably expressing the human TSHR and the Photinus pyralis luciferase gene under transcriptional control of multiple cAMP responsive elements.44 Advantageously, serum was measured in small amounts, and the assay duration time was shorter than for previously described techniques.44,45 Another luminescent-based bioassay employed the CHO-K1 cell line, clone JP09 which was stably transfected with the human TSHR, and the firefly luciferase gene containing cAMP responsive elements.46 The clone “lulu1” was selected by limiting dilution and exhibited dose-dependent TSH response. Furthermore, the assay could be performed on unfractionated serum.46
Chimeric TSHR-Expressing Cell Lines.
To analyze the binding sites for TSH and TSI involved in GD 8 human TSH receptor/rat luteinizing hormone-choriogonadotropin receptor chimera constructs were established.47 Interestingly, 1 of these chimeras, the mutant chimeric four (Mc4) receptor, conserved the ability to interact with TSH and Ab from GD and from idiopathic myxedema patients.47 Ensuing from this work, a reporter gene bioassay was established specifically for the detection of TSI.30 This Food and Drug Administration-cleared reporter bioassay, uses a genetically modified CHO-K1 cell line, stably transformed with a plasmid comprising the firefly luciferase reporter gene under control of the alpha 4 glycoprotein promoter which has cAMP responsive elements and the gene for Mc4 TSHR constitutively expressed by the simian virus 40 promoter. A schematic representation of this TSI bioassay is shown in Figure 3. In the Mc4 TSHR construct, the amino acid residues 262 to 368 of the C-terminus were substituted with the amino acid residues 261 to 329 of the rat luteinizing hormone-choriogonadotropin receptor. The results, obtained within less than 20 hours, were expressed as an activity ratio (specimen-to-reference ratio, SRR %) of patient sample compared with a reference control sample.
Recently, several cell lines were developed after repetitive limiting dilution, mink lung epithelial cells, human embryonic kidney (HEK293) cells, and CHO-K1 cells also expressing the chimeric Mc4 receptor.48,49 In the Mc4 chimera, amino acid residues 261 to 370 from the human TSHR were substituted by the amino acid residues 261 to 329 of the rat LH/CG receptor. Different transfectants were stimulated with forskolin and bTSH, consequently all Mc4 transfected cell lines responded better than the wild-type TSHR cells. The Mc4 responses were not cell type and cAMP responsive element–luciferase construct dependent. As previously shown by others,50 transfected cells with the chimeric Mc4 TSHR were more sensitive and specific than bioassays performed with the wild-type–TSHR. Currently used functional TSHR-Ab bioassays for the measurement of TSI are listed in Table.
ROLE OF TSHR MAb IN AUTOIMMUNE THYROID DISEASE AND TED
Isolation of human TSHR MAb with either stimulating (M22) or blocking activities (K1-70) has been a major advance in studies on the TSHR. The stimulating M22 MAb is widely used in the recently introduced automated TBII binding assay and has been frequently tested in cell biological and immunologic studies dealing with the extrathyroidal expression of the TSHR. M22 enhanced adipogenesis via phosphoinositide 3-kinase activation in orbital preadipocytes from patients with TED51 and increased hyaluronic acid synthesis in Graves orbital fibroblasts.52 A small molecule antagonist that inhibits TSHR-mediated stimulation of cAMP production in Graves orbital fibroblasts very recently demonstrated the key role of the TSHR in the pathogenesis of TED.53 Similarly, K1-70 has been shown to inhibit the thyroid stimulating activity of M22 in vivo.54 Conversely, M22 has been recently applied to compare the analytical performance, the limits of blank, detection and quantitation, and the half maximal effective concentration (EC50) of a Food and Drug Administration-cleared TSI bioassay and an automated binding assay.55 Beyond their significance in the diagnosis and pathogenesis of GD and autoimmune thyroid disease, functional Mab, which act as TSHR antagonists, are potentially important new therapeutic agents for TED. For example, in GD and related TED, K1-70 may well be effective in controlling hyperthyroidism and the eye signs caused by TSI. In addition, hyperthyroidism caused by autonomous TSH secretion, due to TSHR activating mutations,56 should be treatable by K1-70. Finally, K1-70 has potential applications in thyroid imaging and targeted drug delivery to TSHR-expressing tissues.
To assess the clinical relevance of TSI in Graves patients without (GD) or with orbitopathy (TED), to correlate the TSI levels with activity/severity of TED, and to compare the sensitivity/specificity of the TSI bioassay with TSHR-binding methods (TBII), TSI were tested in 2 reporter cell lines designed to measure immunoglobulins binding the TSHR and transmitting signals for cAMP/cAMP response element–binding protein/cAMP responsive element complex-dependent activation of luciferase gene expression.50 All hyperthyroid TED patients were TSI positive. TSHR-stimulating immunoglobulins were detected in 150 of 155 (97%) TED patients and in 0 of 40 controls. Serum TSI titers were 3- and 8-fold higher in TED versus GD and control, respectively. All patients with diplopia, optic neuropathy, and smokers were TSI positive. TSI strongly correlated with clinical activity (r = 0.87 and r = 0.7, both p < 0.001) and clinical severity (r = 0.87 and r = 0.72, both p < 0.001) of TED in the bioassay. Clinical sensitivity (97% vs. 77%, p < 0.001) and specificity (89% vs. 43%, p < 0.001) of the TSI were greater than TBII in TED. All tested TSI-positive/TBII-negative patients had TED, whereas all TSI-negative/TBII-positive subjects had GD only. Thus, based on this study, TSI is a functional indicator of TED activity and severity.
Within a cross-sectional trial, the clinical relevance of these functional Ab was assessed in 108 untreated patients with TED.57 TSI were detected in 106 of 108 (98%) TED patients. All 53 hyperthyroid patients were TSI positive versus 47 (89%) TBII positive. All 69 patients with active TED were TSI positive, whereas only 58 of 69 (84%) were TBII positive. TSI correlated with the activity (r = 0.83, p < 0.001) and severity (r = 0.81, p < 0.001) of TED. All 59 TED patients with diplopia were TSI positive and 50 of 59 (85%) TBII positive. TSI positivity among patients with moderate-to-severe and mild TED were 75 of 75 (100%) and 31 of 33 (94%) compared with TBII positivity of 63 of 75 (84%) and 24 of 33 (73%), respectively. TSI levels were higher in moderate-to-severe versus mild TED (489% ± 137% vs. 251% ± 100%, p < 0.001). Chemosis and TED activity predicted TSI levels alone, p < 0.001 (multivariable analysis). TSI levels were higher in patients with (527% ± 131%) than in patients without chemosis (313% ± 127%, p < 0.001). Thus, based on this study, TSI show more significant association with clinical features of TED than TBII and may be regarded as functional biomarkers for TED.
Within a prospective trial,58 the clinical usefulness of TSI was evaluated in 101 consecutive patients with severe and active TED. At an academic referral multidisciplinary thyroid eye clinic, complete ophthalmic, endocrine, and serologic investigations were performed. All 101 consecutively followed patients with severe and active TED were TSH receptor blocking antibody negative. In contrast, 91 (90%) were TSI positive of whom 90 had GD. Of the 10 TSI-negative patients, 4 had had a thyroidectomy, 2 radioactive iodine treatment (7 and 10 years before), 1 was treated with antithyroid drugs, and 3 were diagnosed with HT. Serum TSI levels correlated with the diplopia score (p = 0.016), total severity eye score (p = 0.009), proptosis (p = 0.007), eyelid aperture (p = 0.003), upper eyelid retraction (p = 0.006), keratopathy (p = 0.04), thyroid binding inhibiting immunoglobulins (TBII, p < 0.001), and negatively with the duration of TED (p = 0.002). Median serum values of TSI and TBII were SRR% 418 (range 28–795%) and 7.35 IU/L (0.3–174 IU/L), respectively. Therefore, based on this study, the stimulating (TSI), not the blocking (TSHR-blocking immunoglobulin), is highly prevalent in severe/active TED, and serum TSI levels correlate with clinical disease severity.
An American retrospective analysis showed also a correlation between the levels of TSI and objective clinical findings in 23 patients with TED.59 The authors reported that changes in the inflammatory phase of TED correlated with changes of the serum TSI levels, thus confirming our own series of more than 100 patients with TED. This interesting report demonstrates that serial serum TSI measurements are an useful tool in assessing clinical inflammatory activity of TED and help management decision-making in patients with TED.
Recognition of dysthyroid optic neuropathy (DON) requires sensitive diagnostic tools. Clinical assessment may fail to reliably evaluate the acuteness of DON especially if signs for inflammation are missing. Aim of a cross-sectional study60 was to assess the relationship between TSI and onset of DON. At a multidisciplinary orbital center, serum TSI levels were measured in 180 consecutive patients with TED and 302 healthy controls. Thirty of 180 (16.7%) patients with TED had DON of recent onset or a history of DON (post-DON). Optic disk swelling was present, and visual-evoked potentials were pathologic in all eyes with DON of recent onset, but in 1 of 13 (7.7%) with post-DON, only (p = 0.005). Nineteen of 20 (96%) patients with DON of recent onset were TSI positive. TSI was associated with DON of recent onset (odds ratio 20.96; 95% confidence interval [CI]: 1.064–412.85, p = 0.045). All controls were TSI negative. TSI correlated with the clinical activity score (r = 0.70, p < 0.001), and higher TS levels were noted in active versus inactive TED (485.1% ± 132.3% vs. 277.7% ± 143.7%, cut off < 140%; p < 0.001). Six of 7 (85.7%) patients with inactive TED with recent onset DON versus 1 of 4 (25%) with active post-DON were TSI positive (p = 0.006). A discriminatory cut point of SRR% 377 for TSI was determined based on a Receiver Operating Characteristic analysis (sensitivity 0.95, specificity 0.80). Thus, based on this study, serum TSI levels identify patients with DON of recent onset requiring urgent therapy.
Because the role of TSH receptor stimulating Ab in pediatric GD was regarded as potentially controversial, a multicenter study evaluated the clinical relevance of TSI in GD children with (TED) and without orbital disease.61 Serum samples from 157 GD children were collected in 7 centers and evaluated in a central laboratory. In 82 untreated GD children, sensitivity, specificity, and positive and negative predictive values for TSI and TBII were 100% and 92.68% (p = 0.031), 100% and 100%, 100% and 100%, 100% and 96.15%, respectively. In all GD children, TSI and TBII were present in 147 of 157 (94%) and 138 (87.89%), p < 0.039 and in 247 of 263 (94%) and 233 (89%) samples, p < 0.0075, respectively. In children with TED, TSI and TBII were noted in 100% and 96%, p < 0.001, respectively. Hyperthyroid TED children showed markedly higher TSI levels compared to those with thyroidal GD only (p < 0.0001). No significant differences were noted for TBII between the two groups. During a median medical treatment of 3 years, the decrease of TSI levels was 69% in GD versus 20% in TED, p < 0.001. All 31 samples of euthyroid TED children were TSI positive; in contrast, only 24 were TBII positive, p = 0.016. All control children were TSI negative. Thus, based on this study, serum TSI levels are a sensitive and specific biomarker for pediatric GD and/or TED and correlate highly with disease severity and extrathyroidal manifestations.
TED rarely occurs in patients with HT. At an academic joint thyroid eye clinic within a longitudinal observational study, the prevalence of TSI in 1,055 HT patients with and without TED and controls was evaluated.62 Of 700 consecutive and unselected patients with HT, 44 (6%) had overt TED. All healthy controls were TSI negative. In contrast, serum was TSI positive in 30 of 44 (68.2%) and 36 of 656 (5.5%, p < 0.001) patients with HT + TED and HT, respectively. Compared to patients with HT only, serum TSI levels were higher in HT + TED (median SRR%, 25% and 75% percentile): 45, 35–65 versus 192.5, 115–455.3, p < 0.001. Highest TSI values were noted in patients with active and severe TED versus those with mild and inactive TED: 486, 392–592 versus 142, 73–192.5, p < 0.001. The odds ratio of TSI positivity for the risk of TED adjusted for gender and age was 55.9 (95% CI: 24.6–127, p < 0.0001), while the odds ratio per 10-fold change in TSI SRR% (quantitative TSI) was 133 (95% CI: 45–390, p < 0.0001). The area under the Receiver Operating Characteristic curve for qualitative and quantitative TSI was 87.2% (95% CI: 80.6–93.8) and 89.4% (95% CI: 84.1–94.7), respectively. Thus, based on this study, TSI is strongly associated with TED in HT, and TSI may contribute to the pathophysiology of TED.
Therefore, several studies have shown the clinical relevance and usefulness of measurement of TSHR-Ab in patients with TED, especially in those with clinically active and severe disease according to the European guidelines for the management of TED63 and GD. Foremost, the functional TSHR-stimulating antibodies or immunoglobulins may be regarded as useful and excellent biomarker for the clinical activity and clinical severity of TED.
EVIDENCE-BASED RECOMMENDATIONS FOR CLINICAL CONTEMPORARY USE
The hyperthyroidism guidelines of the American Thyroid Association64 strongly recommend measurement of TSHR antibodies for the accurate and early diagnosis and management of GD. The conventional competitive-binding (either Enzyme-Linked Immunosorbent Assay or automated) immunoassays for the measurement of TSHR-Ab are available worldwide, and the sensitivity and specificity of the third generation of these binding assays approximate 90–95% and 95–97%, respectively.65 In comparison, the cell-based bioassays are available in the large commercial laboratories in the United States (Quest, Madison, NJ, U.S.A., Labcorp, Burlington, NC, U.S.A. etc.) and in academic reference labs, i.e., Mayo Clinic and Foundation. The bioassays are widely offered in commercial labs in Japan and in selected reference labs in Europe. The costs of ordering these assays depends on several factors i.e., country, state, health insurance, health care provider, etc. In Europe, i.e., at our institution, the costs are identical. When compared one to one with the binding assays, cell-based bioassays are more sensitive (99–100% sensitivity) detecting TSHR-Ab at much lower Ab concentration55,66,67 and are exclusively specific pertaining to differentiation of Ab functionality (100% specificity). In particular, the analytical performance and clinical usefulness of a Food and Drug Administration-cleared, stimulatory TSHR bioassay in patients with GD has been shown.55 In addition, a multicenter trial confirmed the very high specificity, sensitivity, and positive and negative predictive values of this tool for the diagnosis of GD in children.61 Finally, incorporation and early utilization of TSHR stimulatory antibodies in current diagnostic algorithms was shown to confer a 46% shortened time to diagnosis of GD and a cost savings of 47%.68
In conclusion, the measurement of TSHR-Ab in general and functional (especially stimulating) Ab, in particular, is recommended for the rapid diagnosis, differential diagnosis, and management of patients with Graves hyperthyroidism, related TED, and in patients with HT and extrathyroidal manifestations.
1. Kahaly GJ, Olivo PD. Graves’ Disease. N Engl J Med 2017;376:184.
2. Brent GA. Clinical practice. Graves’ disease. N Engl J Med 2008;358:2594–605.
3. Weetman AP. Graves’ disease. N Engl J Med 2000;343:1236–48.
4. Lazarus JH, Marino M. Wiersinga W.M., Kahaly G.J. Graves’ Orbitopathy, in A Multidisciplinary Approach - Question and Answers. 2010: 2nd revised edition., Basel, Switzerland: Karger, 26–32.
5. Kahaly GJ. The thyrocyte-fibrocyte link: closing the loop in the pathogenesis of Graves’ disease? J Clin Endocrinol Metab 2010;95:62–5.
6. Bahn RS. Graves’ ophthalmopathy. N Engl J Med 2010;362:726–38.
7. Hansen C, Fraiture B, Rouhi R, et al. HPLC glycosaminoglycan analysis in patients with Graves’ disease. Clin Sci (Lond) 1997;92:511–7.
8. Hansen C, Rouhi R, Förster G, et al. Increased sulfatation of orbital glycosaminoglycans in Graves’ ophthalmopathy. J Clin Endocrinol Metab 1999;84:1409–13.
9. Kahaly G, Schuler M, Sewell AC, et al. Urinary glycosaminoglycans in Graves’ ophthalmopathy. Clin Endocrinol (Oxf) 1990;33:35–44.
10. Kahaly G, Stover C, Otto E, et al. Glycosaminoglycans in thyroid-associated ophthalmopathy. Autoimmunity 1992;13:81–8.
11. Förster G, Otto E, Hansen C, et al. Analysis of orbital T cells in thyroid-associated ophthalmopathy. Clin Exp Immunol 1998;112:427–34.
12. Kahaly GJ, Shimony O, Gellman YN, et al. Regulatory T-cells in Graves’ orbitopathy: baseline findings and immunomodulation by anti-T lymphocyte globulin. J Clin Endocrinol Metab 2011;96:422–9.
13. Laurberg P, Wallin G, Tallstedt L, et al. TSH-receptor autoimmunity in Graves’ disease after therapy with anti-thyroid drugs, surgery, or radioiodine: a 5-year prospective randomized study. Eur J Endocrinol 2008;158:69–75.
14. Massart C, Gibassier J, d’Herbomez M. Clinical value of M22-based assays for TSH-receptor antibody (TRAb) in the follow-up of antithyroid drug treated Graves’ disease: comparison with the second generation human TRAb assay. Clin Chim Acta 2009;407:62–6.
15. Kahaly GJ, Diana T. TSH receptor antibody functionality and nomenclature. Front Endocrinol (Lausanne) 2017;8:28.
16. Smith BR, Sanders J, Furmaniak J. TSH receptor antibodies. Thyroid 2007;17:923–38.
17. Smith BR, Hall R. Thyroid-stimulating immunoglobulins in Graves’ disease. Lancet 1974;2:427–31.
18. Shewring G, Smith BR. An improved radioreceptor assay for TSH receptor antibodies. Clin Endocrinol (Oxf) 1982;17:409–17.
19. Sanders J, Oda Y, Roberts S, et al. The interaction of TSH receptor autoantibodies with 125I-labelled TSH receptor. J Clin Endocrinol Metab 1999;84:3797–802.
20. Costagliola S, Morgenthaler NG, Hoermann R, et al. Second generation assay for thyrotropin receptor antibodies has superior diagnostic sensitivity for Graves’ disease. J Clin Endocrinol Metab 1999;84:90–7.
21. Bolton J, Sanders J, Oda Y, et al. Measurement of thyroid-stimulating hormone receptor autoantibodies by ELISA. Clin Chem 1999;45:2285–2287.
22. Ando T, Latif R, Davies TF. Thyrotropin receptor antibodies: new insights into their actions and clinical relevance. Best Pract Res Clin Endocrinol Metab 2005;19:33–52.
23. Gassner D, Stock W, Golla R, et al. First automated assay for thyrotropin receptor autoantibodies. Clin Chem Lab Med 2009;47:1091–5.
24. Hermsen D, Broecker-Preuss M, Casati M, et al. Technical evaluation of the first fully automated assay for the detection of TSH receptor autoantibodies. Clin Chim Acta 2009;401:84–9.
25. Schott M, Hermsen D, Broecker-Preuss M, et al. Clinical value of the first automated TSH receptor autoantibody assay for the diagnosis of Graves’ disease (GD): an international multicentre trial. Clin Endocrinol (Oxf) 2009;71:566–73.
26. Kamijo K, Murayama H, Uzu T, et al. Similar clinical performance of a novel chimeric thyroid-stimulating hormone receptor bioassay and an automated thyroid-stimulating hormone receptor binding assay in Graves’ disease. Thyroid 2011;21:1295–9.
27. Rahhal SN, Eugster EA. Thyroid stimulating immunoglobulin is often negative in children with Graves’ disease. J Pediatr Endocrinol Metab 2008;21:1085–8.
28. Botero D, Brown RS. Bioassay of thyrotropin receptor antibodies with Chinese hamster ovary cells transfected with recombinant human thyrotropin receptor: clinical utility in children and adolescents with Graves disease. J Pediatr 1998;132:612–8.
29. Shibayama K, Ohyama Y, Yokota Y, et al. Assays for thyroid-stimulating antibodies and thyrotropin-binding inhibitory immunoglobulins in children with Graves’ disease. Endocr J 2005;52:505–10.
30. Lytton SD, Li Y, Olivo PD, et al. Novel chimeric thyroid-stimulating hormone-receptor bioassay for thyroid-stimulating immunoglobulins. Clin Exp Immunol 2010;162:438–46.
31. Takasu N, Kamijo K, Sato Y, et al. Sensitive thyroid-stimulating antibody assay with high concentrations of polyethylene glycol for the diagnosis of Graves’ disease. Clin Exp Pharmacol Physiol 2004;31:314–19.
32. Uno C, Nishikawa M. [Clinical studies on abnormal thyroid stimulators in patients with Graves’ disease. II. Clinical significance of measuring TSAb and TBII in patients with euthyroid Graves’ disease and patients with hyperthyroid Graves’ disease during antithyroid drug treatment]. Nihon Naibunpi Gakkai Zasshi 1988;64:206–15.
33. Kim MR, Faiman C, Hoogwerf BJ, et al. Thyroid-stimulating antibody assay using a human thyrotropin receptor transfected cell line: relationship to clinical features of graves’ disease. Endocr Pract 1997;3:337–43.
34. Smith BR, Sanders J, Furmaniak J. TSH receptor antibodies. Thyroid, 2007;17:923–38.
35. Rees Smith B, Sanders J, Evans M, et al. TSH receptor - autoantibody interactions. Horm Metab Res 2009;41:448–455
36. Lytton SD, Li Y, Olivo PD, et al. Novel chimeric thyroid-stimulating hormone-receptor bioassay for thyroid-stimulating immunoglobulins. Clin Exp Immunol 2010;162:438–46.
37. Pierce M, Sandrock R, Gillespie G, et al. Measurement of thyroid stimulating immunoglobulins using a novel thyroid stimulating hormone receptor-guanine nucleotide-binding protein, (GNAS) fusion bioassay. Clin Exp Immunol 2012;170:115–21.
38. Araki N, Iida M, Amino N, et al. Rapid bioassay for detection of thyroid-stimulating antibodies using cyclic adenosine monophosphate-gated calcium channel and aequorin. Eur Thyroid J 2015;4:14–9.
39. Parmentier M, Libert F, Maenhaut C, et al. Molecular cloning of the thyrotropin receptor. Science 1989;246:1620–2.
40. Perret J, Ludgate M, Libert F, et al. Stable expression of the human TSH receptor in CHO cells and characterization of differentially expressing clones. Biochem Biophys Res Commun 1990;171:1044–50.
41. Morgenthaler NG, Pampel I, Aust G, et al. Application of a bioassay with CHO cells for the routine detection of stimulating and blocking autoantibodies to the TSH-receptor. Horm Metab Res 1998;30:162–8.
42. Takano T, Sumizaki H, Amino N. Detection of thyroid-stimulating antibody using frozen stocks of Chinese hamster ovary cells transfected with cloned human thyrotropin receptor. Endocr J 1997;44:431–5.
43. Wallaschofski H, Paschke R. Detection of thyroid stimulating (TSAB)- and thyrotropin stimulation blocking (TSBAB) antibodies with CHO cell lines expressing different TSH-receptor numbers. Clin Endocrinol (Oxf) 1999;50:365–72.
44. Watson PF, Ajjan RA, Phipps J, et al. A new chemiluminescent assay for the rapid detection of thyroid stimulating antibodies in Graves’ disease. Clin Endocrinol (Oxf) 1998;49:577–81.
45. Himmler A, Stratowa C, Czernilofsky AP. Functional testing of human dopamine D1 and D5 receptors expressed in stable cAMP-responsive luciferase reporter cell lines. J Recept Res 1993;13:79–94.
46. Evans C, Morgenthaler NG, Lee S, et al. Development of a luminescent bioassay for thyroid stimulating antibodies. J Clin Endocrinol Metab 1999;84:374–7.
47. Tahara K, Ban T, Minegishi T, et al. Immunoglobulins from Graves’ disease patients interact with different sites on TSH receptor/LH-CG receptor chimeras than either TSH or immunoglobulins from idiopathic myxedema patients. Biochem Biophys Res Commun 1991;179:70–7.
48. Giuliani C, Cerrone D, Harii N, et al. A TSHR-LH/CGR chimera that measures functional thyroid-stimulating autoantibodies (TSAb) can predict remission or recurrence in graves’ patients undergoing antithyroid drug (ATD) treatment. J Clin Endocrinol Metab 2012;97:E1080–7.
49. Giuliani C, Cerrone D, Harii N, et al. A TSHr-LH/CGr chimera that measures functional TSAb in Graves’ disease. J Clin Endocrinol Metab 2012;97:E1106–15.
50. Lytton SD, Ponto KA, Kanitz M, et al. A novel thyroid stimulating immunoglobulin bioassay is a functional indicator of activity and severity of Graves’ orbitopathy. J Clin Endocrinol Metab 2010;95:2123–31.
51. Kumar S, Nadeem S, Stan MN, et al. A stimulatory TSH receptor antibody enhances adipogenesis via phosphoinositide 3-kinase activation in orbital preadipocytes from patients with Graves’ ophthalmopathy. J Mol Endocrinol 2011;46:155–63.
52. Kumar S, Iyer S, Bauer H, et al. A stimulatory thyrotropin receptor antibody enhances hyaluronic acid synthesis in graves’ orbital fibroblasts: inhibition by an IGF-I receptor blocking antibody. J Clin Endocrinol Metab 2012;97:1681–7.
53. Neumann S, Pope A, Geras-Raaka E, et al. A drug-like antagonist inhibits thyrotropin receptor-mediated stimulation of cAMP production in Graves’ orbital fibroblasts. Thyroid 2012;22:839–43.
54. Furmaniak J, Sanders J, Young S, et al. In vivo
effects of a human thyroid-stimulating monoclonal autoantibody (M22) and a human thyroid-blocking autoantibody (K1-70). Auto Immun Highlights 2012;3:19–25.
55. Leschik JJ, Diana T, Olivo PD, et al. Analytical performance and clinical utility of a bioassay for thyroid-stimulating immunoglobulins. Am J Clin Pathol 2013;139:192–200.
56. Paschke R, Ludgate M. The thyrotropin receptor in thyroid diseases. N Engl J Med 1997;337:1675–81.
57. Ponto KA, Kanitz M, Olivo PD, et al. Clinical relevance of thyroid-stimulating immunoglobulins in graves’ ophthalmopathy. Ophthalmology 2011;118:2279–85.
58. Kampmann E, Diana T, Kanitz M, et al. Thyroid stimulating but not blocking autoantibodies are highly prevalent in severe and active thyroid-associated orbitopathy: a prospective study. Int J Endocrinol 2015;2015:678194.
59. Dragan LR, Seiff SR, Lee DC. Longitudinal correlation of thyroid-stimulating immunoglobulin with clinical activity of disease in thyroid-associated orbitopathy. Ophthal Plast Reconstr Surg 2006;22:13–9.
60. Ponto KA, Diana T, Binder H, et al. Thyroid-stimulating immunoglobulins indicate the onset of dysthyroid optic neuropathy. J Endocrinol Invest 2015;38:769–77.
61. Diana T, Brown RS, Bossowski A, et al. Clinical relevance of thyroid-stimulating autoantibodies in pediatric graves’ disease-a multicenter study. J Clin Endocrinol Metab 2014;99:1648–55.
62. Kahaly GJ, Diana T, Glang J, et al. Thyroid stimulating antibodies are highly prevalent in Hashimoto’s thyroiditis and associated orbitopathy. J Clin Endocrinol Metab 2016;101:1998–2004.
63. Bartalena L, Baldeschi L, Boboridis K, et al; European Group on Graves’ Orbitopathy (EUGOGO). The 2016 European Thyroid Association/European Group on Graves’ orbitopathy guidelines for the management of graves’ orbitopathy. Eur Thyroid J 2016;5:9–26.
64. Ross DS, Burch HB, Cooper DS, et al. 2016 American Thyroid Association Guidelines for diagnosis and management of hyperthyroidism and other causes of thyrotoxicosis. Thyroid 2016;26:1343–421.
65. Tozzoli R, Bagnasco M, Giavarina D, et al. TSH receptor autoantibody immunoassay in patients with Graves’ disease: improvement of diagnostic accuracy over different generations of methods. Systematic review and meta-analysis. Autoimmun Rev 2012;12:107–13.
66. Diana T, Wüster C, Kanitz M, et al. Highly variable sensitivity of five binding and two bio-assays for TSH-receptor antibodies. J Endocrinol Invest 2016;39:1159–65.
67. Diana T, Wüster C, Olivo PD, et al. Performance and specificity of 6 immunoassays for TSH receptor antibodies: a multicenter study. Eur Thyroid J 2017;6:243–9.
© 2018 by The American Society of Ophthalmic Plastic and Reconstructive Surgery, Inc., All rights reserved.
68. McKee A, Peyerl F. TSI assay utilization: impact on costs of Graves’ hyperthyroidism diagnosis. Am J Manag Care 2012;18:e1–14.