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Original Studies |
I.R.I.B.H.N., ULB (S.C., J.E.D., G.V.), and Euroscreen (S.C.), Brussels, Belgium; and Research Laboratories, B.R.A.H.M.S Diagnostica GmbH (N.G.M., J.S., W.W., J.M.H., A.B.), D-12099 Berlin; the Division of Endocrinology, Department of Medicine, University of Essen (R.H., S.P., B.Q., K.M.), D-45122 Essen; and the Center of Internal Medicine, University of Frankfurt (K.B., D.F., P.-M.S.-D., K.-H.U.), D-60950 Frankfurt, Germany
Address all correspondence and requests for reprints to: Dr. Nils G. Morgenthaler, Research Laboratories, B.R.A.H.M.S Diagnostica GmbH, Komturstrasse 19–20, D-12099 Berlin, Germany. E-mail: morgenthaler{at}brahms.de or Prof. Dr. Klaus-Henning Usadel, Zentrum
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Abstract |
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 Top Abstract IntroductionSubjects and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionSubjects and MethodsResultsDiscussionReferences |
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Other detection systems have been described, such as autoantibody detection by FACS (11, 12), immunoprecipitation (13), immunocytochemistry (14), or transferring serum to nude mice (15). These methods are still in an experimental state and are too time consuming and cumbersome for everyday use.
The only validated routine assays for the detection of TRAb are RRAs using porcine thyroid membrane extracts based on the method of Shewring and Rees Smith (16). In their commercialized form, these assays are widely used. Although about 70–90% of TRAb in the serum of Graves’ disease patients are detected by these RRA, there is evidence of clinical hyperthyroid patients classified as Graves’ disease who are negative in the RRA using porcine-derived antigen (17, 18). To increase the sensitivity of the assay, many groups have tried to replace the porcine source of TSH receptor by human recombinant antigen. This approach was reported to work on a small scale (19, 20), but the difficulties associated with the large scale production of human TSH receptor have limited its applicability in clinical practice. These early experiments were performed with mammalian cell lines, which produced a well glycosylated TSH-R that binds TSH and autoantibodies efficiently. However, the growth conditions of these cells and the level of receptor expression did not permit the production of sufficient antigen for routine assays. Alternative approaches by overexpressing TSH-R in bacteria (21, 22) or insect cells (23, 24, 25) did not lead to the expression of a functional receptor, i.e. binding both TSH and autoantibodies.
Here we show that expression of the human TSH-R (hTSH-R) in the leukemia cell line K562 yielded adequate amounts of bioactive human antigen. In addition, a monoclonal antibody (moAb) (26) recognizing only the native receptor has allowed for the development of a new RRA in the coated tube (CT) format as well as a nonradioactive chemiluminescence assay. The clinical superiority of these assays compared to the conventional RRA was demonstrated with 328 sera from patients with Graves’ disease and 520 different controls.
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Subjects and Methods |
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TopAbstractIntroductionSubjects and MethodsResultsDiscussionReferences |
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Included in the study were 328 patients with Graves’ disease. Graves’ disease was diagnosed initially according to standard clinical criteria (suppressed TSH, elevated T3 or free T4, goiter, sonography, and signs of Graves’ ophthalmopathy when present). Patients were grouped according to their metabolic state in patients with active hyperthyroid Graves’ disease without or with less than 4 weeks of treatment with antithyroid drugs (group 1, n = 86), patients receiving treatment for longer than 4 weeks (group 2, n = 126), and patients in remission without treatment (group 3, n = 116).
Furthermore, we included 54 patients with Hashimoto’s thyroiditis
(diagnosed on the bases of clinical hypothyroidism, sonography, and the
presence of antithyroid peroxidase and/or antithyroglobulin
autoantibodies; group 4), 69 patients with nonthyroid autoimmune
diseases (type I diabetes, rheumatoid arthritis, and systemic lupus
erythematosus) with nonthyroid autoantibodies (group 5), 115 patients
with goiter but no signs of autoimmune thyroid disease (group 6), and
282 healthy individuals without a history of thyroid disease who were
euthyroid and negative for antithyroid peroxidase and antithyroglobulin
autoantibodies (group 7). The clinical data of the patients are
summarized in Table 1
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Generation of recombinant TSH-R-producing cells
The coding sequence of the hTSHR complementary DNA (27) was subcloned into the KpnI/XbaI sites of pEFIN, a bicistronic vector developed at EUROSCREEN (Brussels, Belgium) (28). K562 cells were maintained at 37 C and 5% CO2 in DMEM containing 10% FCS. For transfection, 2 x 106 exponentially growing cells were electroporated at 0.2 kV/cm, 960 µF (Bio Rad Gene Pulser) for 25 ms in the presence of 20 µg specific linearized plasmid DNA and seeded into culture flask. Forty-eight hours after electroporation, selection was started with 800 µg/mL G418 (Life Technologies, Grand Island, NY). After cloning by limiting dilution, a stable clone expressing high levels of hTSHR was selected. The number of receptors expressed per cell and their dissociation constant (Kd) were computed from displacement curves in which binding of 125I-labeled bovine TSH (bTSH) was competed for by increasing concentrations of unlabeled bTSH. It was estimated that the cells harbored about 1 x 106 receptors with a Kd of 4.8 mIU/mL.
Selection of moAb by FACS
K562 cells expressing hTSHR or K562 cells (wild type) were
transferred into Falcon 2052 tubes (200,000 cells/tube;
Becton Dickinson, Mountain View, CA). Cells were
centrifuged at 500 x g at 4 C for 3 min, and the
supernatant was removed by inversion. They were incubated for 30 min at
room temperature with 100 µL phosphate-buffered saline (PBS)-0.1%
BSA containing 10 µL culture supernatant from 3 different hybridomas
(BA8, 3G4, 5A6) (26). The cells were washed with 4 mL PBS-0.1% BSA and
centrifuged as above. They were incubated for 30 min on ice in the dark
with fluorescein-conjugated 
-chain-specific goat antimouse IgG
(Sigma Chemical Co., St. Louis, MO) in the same buffer.
Propidium iodide (10 µg/mL) was used for detection of damaged cells
that were excluded from the analysis. Cells were washed once again and
resuspended in 250 µL PBS-0.1% BSA. The fluorescence of 5,000
cells/tube was assayed by a FACScan flow cytofluorometer (Becton Dickinson and Co., Eerenbodegem, Belgium). The murine moAb BA8
bound specifically to the native human TSH-R (26) and was used for
subsequent studies.
Preparation of hTSH-R
K562 cells expressing the hTSH-R (K562-TSH-R) were grown in Spinner flasks at 37 C and 5% CO2 in DMEM containing 10% FCS (without G418) to a density of 1.5 x 106 cells/mL. Cells were harvested by centrifugation (2,400 x g, 10 min, 4 C), washed once with ice-cold PBS, and stored at -80 C. Frozen cells were resuspended in washing buffer [50 mmol/L HEPES (pH 6.8) and 50 mmol/L NaCl] and centrifuged (100,000 x g, 30 min, 4 C). The cell pellet was resuspended in extraction buffer [100 mmol/L HEPES (pH 6.8), 2% Triton X-100, and Complete protease inhibitors (Boehringer Mannheim, Mannheim, Germany)] and homogenized in a Potter homogenizer (Braun, Melsungen, Germany). After centrifugation (100,000 x g, 30 min, 4 C), BSA (protease free, Sigma Chemical Co.) was added to a final concentration of 1% to the supernatant containing the solubilized TSH-R.
Production of TSH-R coated tubes
The moAb BA8 (26) was coated for 20 h on polystyrene tubes (1.5 µg/tube) in 0.3 mL buffer [10 mmol/L Tris-HCl (pH 7.5) and 50 mmol/L NaCl]. Tubes were blocked with 10 mmol/L sodium phosphate buffer containing 3% Karion FP and 0.5% protease-free BSA (Sigma Chemical Co.), pH 6.8. The TSH-R-containing extract was diluted 1:50 and added to the tubes. Affinity binding of the TSH-R to the antibody was performed at 4 C for 20 h. Tubes were blocked again [50 mmol/L HEPES (pH 6.5), 0.25% Triton X-100, 1% Karion FP, and 0.5% BSA] and lyophilized. Ten liters of culture medium resulted in the preparation of 20,000 coated tubes.
Preparation of labeled bTSH
bTSH was affinity purified from bovine pituitaries (final activity, 50–60 TSH IU/mg protein) and labeled with 125I using the chloramine-T method, yielding a specific activity of 58 µCi/µg protein. Acridinium ester-labeled bTSH was produced as follows. bTSH (100 µg; 50–60 TSH IU/mg protein) in 20 mmol/L sodium phosphate buffer, pH 7.0, was incubated for 15 min at room temperature with 10 µL acridinium ester (1 mg/mL in acetonitril; Hoechst AG, Frankfurt, Germany). Labeled bTSH was purified by high performance liquid chromatography using a Waters-Protein Pak SW 125 column (running buffer, 0.1 mol/L ammonium acetate, pH 5.5; flow rate, 0.6 mL/min).
Autoantibody measurement in conventional TBII assay
Autoantibody measurement was performed with a commercial RRA (TRAK-Assay, B.R.A.H.M.S Diagnostica) following the manufacturer’s instructions. The functional assay sensitivity of this RRA is 8 U/L.
Autoantibody measurement in CT TBII assays
Patients’ samples or standards (100 µL) were added in
duplicate to hTSH-R coated tubes. To this were added 200 µL buffer
containing 100 mmol/L HEPES, 20 mmol/L ethylenediamine tetraacetate,
0.5 mmol/L N-ethyl-maleimide, 0.1 mmol/L leupeptin, 1% BSA,
0,5% Triton X-100, and 5 µg antihuman TSH antibody (Sigma Chemical Co.), pH 7.5. After 2-h incubation under shaking (300
U/min) at room temperature, tubes were washed once with 2 mL washing
buffer. Then, 200 µL tracer were added containing either
125I- or acridinium ester-labeled bTSH (1 ng/tube;
B.R.A.H.M.S Diagnostica), followed by 1-h incubation at room
temperature. Tubes were washed twice with 2 mL washing buffer, and
detection was performed in either a -counter or a luminometer.
Calibration of CT assays
To compare data between individual test runs, all raw data [counts per min for radioligand assay, relative light units (rlu) for chemiluminescence assay] were expressed in TRAK units as calculated from a standard curve that was included in every run. To obtain a representative standard curve, 20 sera from Graves’ disease patients with high titers were pooled, diluted, and calibrated using the TRAK-Assay.
The Medical Research Council standard of 1966 (long acting thyroid stimulator, lot 65/122) and the WHO standard of 1995 (TSAb, lot 90/672) were tested in both assays. One Medical Research Council unit resulted in approximately 1.5 TRAK units, and 1 WHO unit resulted in 4 TRAK units, respectively.
Definition of cut-off and statistical analysis
To obtain the optimal decision threshold level for positivity,
receiver operating characteristic (ROC) analysis was performed (29).
Sensitivity/specificity pairs were calculated by varying the decision
threshold levels over the entire range of TRAK units. The sensitivity
(true positive results) of all three assays was calculated from the 86
patients in group 1 (untreated Graves’ disease). On the other side,
the specificity (true negative results) was calculated from 282 healthy
blood donors (group 7). The experimental cut-off was determined for all
assays at 99.6% specificity. Statistical analysis was performed using
2 test with Yates correction for comparison of the
autoantibody prevalence between the different assays within the
respective groups and Mann-Whitney rank sum analysis for comparison of
the autoantibody levels in the different groups determined with one
assay. Correlation analysis was performed with Pearson correlation. For
method comparison of CT RRA and TRAK assay, the nonparametric approach
according to Passing and Bablok was used (30).
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Results |
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TopAbstractIntroductionSubjects and MethodsResultsDiscussionReferences |
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). Competition binding experiments with
a [125I]bTSH tracer indicated that the cell line
expresses about 106 receptors/cell with a
Kd of 4.8 mIU/mL (Fig. 1A).
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shows
representative [125I]TSH binding in the presence of
different sera and unlabeled TSH. Similar results were obtained using
the acridinium ester-labeled bTSH in chemiluminescence detection (data
not shown).
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shows
individual sera and a representative standard curve of pooled sera for
the CT LRA and CT RRA. Measuring the Medical Research Council long
acting thyroid stimulator standard (see Materials and
Methods) and the WHO TSAb standard, 1 Medical Research Council U/L
resulted in 1.5 TRAK U/L, and 1 WHO U/L resulted in 4 TRAK U/L,
respectively, in both assays.
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for the CT
RRA. The FAS for the CT LRA has the same value (data not shown).
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shows a plot of the sensitivity and specificity of all three assays.
With a specificity of 99.6% at a cut-off of 4 U/L, the sensitivity of
both CT assays was 98.8% compared to 99.6% specificity and 80.2%
sensitivity of the conventional system at a cut-off of 11 U/L. Using
this cut-off for all groups of patients the sensitivity of the new
assays compared to that of the conventional assay is shown in Table 2. In all three groups of patients with
Graves’ disease (groups 1–3), the CT assays were significantly more
sensitive for the disease than the conventional assay (by
2 test with Yates correction: group 1, 2
= 14.0, P < 0.001; group 2, 2 = 4.1,
P < 0.05; group 3, 2 = 4.0,
P < 0.05). In contrast, there was no significant
difference between the assays in the control groups 5, 6, and 7.
Although the CT assays detected about 10% more patients in group 4
(Hashimoto’s thyroiditis) than the conventional RRA, the difference
was not significant (2 = 1.62; P =
0.23).
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for all
three assays. In all three assays there were significantly higher
autoantibody levels in the groups with Graves‘ disease than in all
control groups, including group 4 (P < 0.0001, by
Mann-Whitney rank sum test). Group 1 (untreated Graves’ disease) and
group 2 (treated Graves’ disease) showed no difference in the
conventional RRA (P = 0.65), although there was a
stronger (but still not significant) difference in the CT assays
between those groups (P = 0.08). However, in all three
assays, groups 1 and 2 had significantly higher autoantibody levels
than group 3 (Graves’ patients in remission; P <
0.0001). Neither assay showed a significant difference in the
autoantibody levels in the healthy controls (group 7), patients with
nonthyroid autoimmunity (group 5), or patients with nonautoimmune
thyroid disease (group 6), but all three assays showed a significant
difference between the Hashimoto’s thyroiditis patients (group 4) and
the healthy controls (P < 0.0001). Although patients
and controls were from two centers, there was no heterogeneity,
i.e. no differences within the groups between the two
centers were observed.
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(r =
0.91; P < 0.001).
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Discussion |
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TopAbstractIntroductionSubjects and MethodsResultsDiscussionReferences |
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Routine use of recombinant hTSH-R in TBII assays has not been practical until now because the high yield production systems, like bacteria (21, 22) or baculovirus (23, 24, 25), did not produce bioactive receptor (i.e. showing TSH binding). The available CHO cell lines (7, 11, 12, 33) do produce bioactive receptor, but their growth conditions for large scale production are fastidious. The K562 line described in the present study provides an efficient solution to this problem; it grows in suspension to densities of 1.5 x 106 cells/mL and expresses stably about 106 receptors/cell.
Our results demonstrate a close to 100% sensitivity of the second generation TBII assay in hyperthyroid patients with Graves’ disease. Thus, this assay using recombinant hTSH-R is clearly superior to its predecessor based on porcine thyroid membrane preparations. The improved sensitivity for Graves’ disease from 80% to nearly 100% may be due to several properties. First, there may be a better configurational accessibility of the recombinant hTSH-R preparation for its antibodies that is not present on the crude porcine membrane preparation. Second, solid phase technology allows for a reduction in nonspecific binding. The better signal to noise ratio leads to a lower decision threshold for the cut-off and increases the sensitivity, as shown by ROC plot analysis. For clinical purposes this high sensitivity allows a rapid distinction between autoimmune and other forms of hyperthyroidism and could obviate the need for other diagnostic procedures in thyrotoxic patients positive for TRAb. Thus, the new assay represents an improvement in the management of Graves’ disease.
Currently, TRAb are measured at the beginning and during the course of antithyroid drug treatment to detect early relapses. A large multicenter study (34), among others, has confirmed that patients with elevated TRAb at the end of drug treatment have a significantly higher relapse rate than those without. However, the low sensitivity and specificity of the assay precluded its use in the prediction of the individual clinical course (34). A meta-analysis combining studies of relapse prediction in Graves’ disease later showed again that TRAb activity is clearly associated with relapse. However, a considerable proportion of TRAb-positive or -negative patients was found in the remission or relapse groups, respectively (35). These studies were based on porcine membrane preparations; consequently, a prospective evaluation of the new assay is warranted that evaluates its predictive value for relapse or remission in the long term course of Graves’ disease.
The median levels of TRAb titers were significantly higher in patients with active Graves’ disease with or without antithyroid drug treatment than in those in remission, although the latter still contained about 48% TRAb positives (compared with 35% of TRAb-positive patients in the porcine membrane assay). Similarly, 15% of patients with Hashimoto’s thyroiditis had detectable TRAb titers using the new CT assays in contrast to 6% with the conventional assay. These TRAb levels, although measurable, are functionally different, as they are not associated with clinical or biochemical hyperthyroidism. They may have binding and blocking capabilities in contrast to the stimulating Igs found in active Graves’ disease. Autoantibodies that block the binding of TSH to its receptor may even lead to hypothyroidism, as in certain cases of atrophic thyroiditis (primary myxedema). Such functional differences can only be distinguished when comparing the binding inhibition assay with the measurement of stimulating Igs (4, 5, 6, 7, 8, 9, 10).
Low TRAb levels were also detected in a limited number of individuals belonging to the control groups (1 of 282 healthy controls, 3 of 115 in nonautoimmune thyroid disease patients, and two out of 69 with nonthyroid autoimmune disease). The significance of these positive values remains to be determined. A close follow-up of these individuals will reveal whether the new TBII assays will be useful for the early detection of autoimmune hyperthyroidism.
Although quantitative correlation between TSAb and TBII assays is relatively poor (10), only a very small number of patients who are clearly TSAb-positive score negative in current TBII assays (10). This, which constitutes the main justification for the routine clinical use of TBII assays, may indicate that stimulation of the receptor by TSAbs implies recognition of an epitope(s) that is part of the TSH-binding site; alternatively, it is compatible with the coexistence in Graves’ patients of stimulating antibodies (that may not always be endowed with TBII properties) and antibodies that compete for TSH binding (and may not be endowed with TSAb or TBAb properties) (36).
Identification of the epitopes corresponding to the various autoantibody categories is a major focus of current research (37). The view that TSAbs and TBAbs would recognize distant parts of the TSH-R ectodomain (36) will need experimental confirmation by the isolation of the individual TRAbs from patient’s serum. Similarly, the respective importance of the nature of autoantibodies or intrinsic/paracrine thyroid factors to explain the variability in goiter size in Graves’ patients remains to be defined.
It is hoped that future generations of TRAb assays will allow for the measurement of the spectrum of autoantibodies with specific functional characteristics. In the meantime, with their increased sensitivity, we consider that the new TBII assays described here constitute a significant improvement over the currently available assays and deserve evaluation in additional prospective studies.
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Acknowledgments |
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Footnotes |
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Received August 6, 1998.
Revised October 6, 1998.
Accepted October 13, 1998.
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References |
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