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Department of Microbiology and Immunology, College of Medicine, University of Illinois at Chicago, Chicago, IL 60612; and
Division of Reproductive Biology, Department of Gynecology/Obstetrics, Stanford University School of Medicine, Stanford, CA 94305
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Since cloning of TSHR, many laboratories have attempted to develop an animal model for GD. Our earlier attempts to induce GD using thyroid membranes as a source of native TSHR was unsuccessful. This, perhaps, was due to very low levels of TSHR expressed on thyroid membranes, thus resulting in inoculation of suboptimal doses of Ag. To overcome this limitation, we expressed large quantities of nonglycosylated and glycosylated extracellular domains of TSHR (ETSHR and hETGP, respectively) in insect cells and showed that they bind TSH and patient autoantibody, respectively (3, 4). We used the ETSHR protein to immunize different strains of mice and showed that only BALB/c mice developed mild hyperthyroidism (5). Subsequently, to enhance the severity of the disease, we primed BALB/c mice with ETSHR and then challenged them with soluble thyroid membranes (5). These mice showed significant elevation in Abs against native TSHR, with concomitant increase in tetraiodothyronine (T4) hormone levels. However, hormonal levels returned to normal within a relatively short time after cessation of Ag administration, suggesting that tolerance to endogenous TSHR had not been overcome. Similarly, a number of different laboratories have attempted to induce GD in mice using various preparations of TSHR, but with only limited success (6, 7, 8, 9, 10, 11, 12). Inability to induce GD using only recombinant TSHR proteins is most likely due to inappropriate folding of the protein. Thus, availability of large quantities of TSHR with native conformation remained a major impediment to developing an appropriate animal model.
To overcome this, Shimojo et al. used RT4.15HP cells, doubly transfected with cDNAs encoding human TSHR and MHC class-II molecules, to immunize AKR mice (13). Approximately 20% of mice immunized with cells expressing both TSHR and class II, but not with cells expressing either class II or TSHR alone, developed hyperthyroidism. Based on these results, the authors suggested that coexpression of MHC class II and TSHR is essential to induce GD. Recently, Kita et al. (14) have confirmed these findings using similarly transfected RT4.15HP cells.
To optimize conditions for disease induction in a higher proportion of mice, we compared the efficacy of full-length TSHR vs soluble ectodomain; mouse TSHR (mTSHR) vs human TSHR (hTSHR); soluble purified protein (TBP) vs cell associated protein (293-TBP cells); and syngeneic (M12 cells) vs xenogeneic cells (293 cells) expressing TSHR. Our current studies clearly demonstrate that immunization of BALB/c mice using either the ectodomain of TSHR with native conformation (TBP), or full-length TSHR protein expressed on cell surface leads to disease induction in almost 100% of mice, with significant elevation in stimulatory Abs and thyroid hormones in their circulation.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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M12 cells were transfected with pSR
puro vector in serum-free
RPMI 1640 containing either a cDNA-encoding hTSHR (hM12 cells) or cDNA
encoding mTSHR (mM12 cells). The pSR vector (15)
consists of a Simian virus (SV40) early promoter and part of the R-U5
segment of the long terminal repeat (LTR) from human T cell leukemia
virus type I. M12 cells were transfected with the plasmids using
Lipofectamine (Life Technologies, Gaithersburg, MD) following the
manufacturer’s protocol and cultured in RPMI 1640 medium. Transfected
cells were selected for puromycin resistance. TSHR expression on stable
transfectants was confirmed by flow cytometry using TSHR-specific Abs
and radiolabeled bovine TSH (bTSH) binding, and by their ability to
produce cAMP in the presence of bTSH. These permanently transfected
hM12 and mM12 cells were used throughout the study.
Flow cytometry to detect TSHR expression
Human embryonic kidney cells (293 cells) expressing extracellular domain of hTSHR (293-TBP cells) (16, 17) and hM12 and mM12 cells, and untransfected control cells were detached from culture plates using PBS-based enzyme-free dissociation solution (Nozyme, Speciality Media, Lavallette, NJ) and transferred into tubes as described earlier (18). Cells were pelletted at 500 x g at 4°C for 5 min, and washed once in PBS containing 1% BSA and 0.1% sodium azide (FACS buffer). Cells were incubated for 30 min at 4°C with 100 µl FACS buffer containing either 1 µl of control rabbit serum, polyclonal rabbit anti-hTSHR, or anti-mTSHR serum (10). Cells were washed three times with 1 ml FACS buffer and centrifuged as above. Cells were subsequently incubated for 30 min on ice in dark with fluorescein-conjugated goat anti-rabbit IgG, then washed three times and suspended in 400 µl of FACS buffer. Propidium iodide (10 µg/ml) was used to detect dead cells. Cells were analyzed using a flow cytometer (Coulter, Hialeah, CA).
cAMP assay
Control M12, hM12, and mM12 cells were plated into 96-well plates in RPMI 1640 supplemented with 10% FBS. Fresh medium (RPMI 1640) was added to these wells 24 h later. When cells grew to confluency, medium was removed, and cells were washed once using RPMI 1640 medium. To these wells, a serial dilution of bTSH in a hypotonic medium (HBSS) containing 0.5 mM 3-isobutyl-methyl-xanthine was added and incubated for 3 h at 37°C. Culture supernatants were collected, and the cAMP released into the medium was measured by RIA (1:100 dilution) using a commercial kit (NEN Life Science Products Incorporated, Boston, MA) following the manufacturer’s protocol.
To test the ability of mouse sera to stimulate cAMP production, Chinese hamster ovary (CHO) cells expressing hTSHR were used (19, 20). Cells were grown to confluency in 96-well plates in F-12 medium supplemented with 10% FBS. Before the assay, medium was removed from the wells, and appropriately diluted sera in hypotonic HBSS containing 0.5 mM 3-isobutyl-ethyl-xanthine were added and incubated for 3 h in a CO2 incubator at 37°C. Culture supernatants were collected and assayed for cAMP production as described above.
TBII assay to detect TSHR-specific Abs
A commercial radioreceptor (Kronus, Dana Point, CA) assay is routinely used to measure autoantibodies to TSHR in the sera of patients with thyroid disorders. In this assay, the ability of Abs to bind to TSHR on porcine thyroid membranes and block the binding of 125I-labeled TSH is measured (i.e., thyrotropin-binding inhibitory index) (TBII)). This assay was used to detect autoantibodies to TSHR according to the manufacturer’s protocol, and results were expressed as TBII activity.
Removal of TBII activity by hM12 and mM12 cells
Cells (M12, hM12, and mM12) were grown to confluency in RPMI 1640 medium supplemented with 10% FBS in a 24-well plate. Before assay, cells were washed three times with buffer containing 10 mM Tris-HCL (pH 7.4), 50 mM sodium chloride, and 0.1% BSA. Fifty microliters of Graves’ patient sera was added to triplicate wells. After incubation for 2 h at room temperature, sera were collected and tested for their TBII activity as described above. Ability of transfected cells to react with anti-TSHR Ab and adsorb the TBII activity of the sera was calculated and reported as percentage blocking.
TSH binding assay
M12, mM12, and hM12 cells were grown to confluency in RPMI 1640, and 293 and 293-TBP cells (16, 17) were grown in DMEM-F12 supplemented with 10% FBS in a 24-well plate (0.5 x 106 cells). On the day of assay, cells were washed three times with buffer containing 10 mM Tris-HCL (pH 7.4), 50 mM sodium chloride, and 0.1% BSA. To each well, 100 µl of (8000 cpm) 125I-labeled bTSH (Kronus) was added and incubated for 2 h at room temperature. Cells were washed with ice-cold buffer, and then 500 µl of 3 N sodium hydroxide was added to lyse the cells. Contents of the wells were carefully transferred to individual test tubes and counted in a gamma counter to detect 125I. Results are expressed as cpm bound.
Purification of TBP protein
293-TBP cells (16, 17) were grown in DMEM/F12 with
10% FBS and 200 µg/ml of G418. Soluble TBP was purified using a
previously described procedure (17). Briefly, the
ectodomain of hTSHR (aa 1–390) was fused to transmembrane and
cytoplasmic regions of CD8 through aa 36–66 of thrombin receptor. To
express the ectodomain of human TSHR in human embryonic kidney 293
cells, the chimeric cDNA was subcloned into pcDNA3 (Invitrogen, San
Diego, CA) and transfected into 293 cells and grown in DMEM/F12 with
10% FBS. 293-TBP cells were grown to confluency in T-150 tissue
culture flasks. The medium was replenished with 20 ml serum-free
DMEM/F12 containing 300 U -thrombin (Enzyme Research Laboratories,
South Bend, IN) and incubated overnight at 37°C. Medium containing
released TBP was collected, centrifuged, and incubated at 4°C with
Sepharose Fast-flow resin covalently linked with nickel. The protein
was eluted with 200 mM imidazole. The eluant was concentrated using a
centricon-30 (Amicon, Bedford, MA) and stored at -70°C until use.
Unlike ectodomains produced in insect cells, the TBP produced in 293
cells is readily soluble.
Animals
Six- to 8-wk-old female BALB/c mice were obtained from The Jackson Laboratory, Bar Harbor, ME. Experiments were approved and conducted according to established institutional review board guidelines.
Immunization of BALB/c mice with M12 cells expressing TSHR
BALB/c mice (five mice per group) were immunized i.p. six times
as per the following schedule: group 1, 2 x 10
7 M12 cells (1); group 2, hM12 cells
along with cholera toxin B subunit (CTB) (2); group 3,
mM12 cells along with CTB (3); group 4, mice primed once
with hETGP and then challenged with hM12 along with CTB
(4); group 5, mice primed once with mETGP (glycosylated
ectodomain of mTSHR expressed in insect cells) and then challenged with
mM12 cells along with CTB (5); group 6, mM12 cells alone
(6); and group 7, hM12 cells alone (7).
Before immunization, the cells were treated with mitomycin C. Blood was
collected from these mice on days indicated in the legends to Figs. 4
and 5, and serum was tested for Abs and thyroid hormones.
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BALB/c mice were immunized i.p. six times as per the following
schedule: group 1, 2 x 107, 293 cells;
group 2, purified TBP protein given along with CTB; group 3, primed
once with purified TBP emulsified in CFA and then challenged with
293-TBP cells with CTB; group 4, 293-TBP cells along with CTB; and
group 5, primed once with hETGP in CFA and then challenged with 293-TBP
cells along with CTB. Before inoculation, cells were treated with
mitomycin C. Blood was collected periodically on days indicated in the
legends to Figs. 4
and 5 and was tested for autoantibodies and thyroid
hormone levels.
ELISA to detect mouse Ig binding to recombinant TBP protein
ELISA plates were coated with soluble TBP protein overnight at 4°C .The antisera obtained from different groups of mice were diluted serially and assayed by ELISA as described earlier (21).
Measurement of total T4 and T3
Total T4 and triiodothyronine (T3) in serum samples were determined using a commercially available assay kit, Coat-A-Count (Diagnostic Products, Los Angeles, CA). Briefly, test tubes coated with anti-T4 or anti-T3 Abs were incubated with sera and 1 ml of 125I-labeled T4/T3 for 1 h at 37°C. The contents of the tubes were decanted, and radioactivity bound to the tubes was quantitated in a gamma counter. The amount of T4 or T3 in the test sample was calculated by plotting the results against a reference curve generated using standards supplied with the kit.
Thyroid histology
Mouse thyroids were removed and placed in 5 ml of 10% formalin. Tissues were embedded in paraffin, and 5-micron thick sections were prepared and stained with hematoxylin and eosin. Multiple sections from each thyroid were examined.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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To test the ability of full-length mTSHR and hTSHR proteins to
induce GD in mice, we developed permanently transfected M12 cells
(21), a B lymphoblastoid cell line derived from BALB/c,
expressing either mTSHR (mM12) or hTSHR (hM12). As shown in Fig. 1, A and B, we
confirmed the expression of both mTSHR and hTSHR proteins by flow
cytometry. We further confirmed TSHR expression by showing TSH binding
to transfected cells but not to control M12 cells (Fig. 2). Moreover, these cells produced cAMP
in response to stimulation by bTSH (Fig. 3). The antigenic nature of the expressed
protein was confirmed by their ability to neutralize autoantibodies in
the sera of patients with GD (Table I).
Preincubation of Graves’ sera with hM12 or mM12 cells almost
completely neutralized the Abs (reduction of TBII by 98% and 85%,
respectively). Together, these experiments clearly demonstrated that
M12 cells were expressing functional TSHRs.
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and Table II, we immunized seven groups of mice
with various permutations and combinations as described under
Materials and Methods. Mice immunized with and without CTB,
or primed with mETGP/hETGP protein, before immunization with hM12 or
mM12 cells, respectively, developed high titers of Abs as measured in
an ELISA (Fig. 4A) with concomitant increase in the TBII
activity (Fig. 4B). Subsequently, we tested their sera for
circulating thyroid stimulatory Abs. As shown in Fig. 4C,
sera from mice in the experimental groups, and not from the control
group, were able to induce high levels of cAMP production (mean ranging
from 6.52 to 20.83 pmol/ml relative to 1.83 pmol/ml for controls, Table II) in CHO cells permanently transfected with a full-length human
TSHR.
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D). Serum T3 levels were also significantly elevated in all
the experimental groups, but not in the control group (mean of
156.18–195.13 ng/dl, for test groups vs 34.99 ng/dl for controls) as
shown in Fig. 4E. Moreover, mice in several experimental
groups showed a tendency toward lower body weight, with mice in group 4
showing significantly lower weights (Fig. 4F), than the
control mice (mean body weight of 19.5–23.24 g for the test group vs
23.9 g for controls). Together, these studies clearly showed that
mice immunized with TSHR had developed pathogenic Ab responses to TSHR
and showed perturbation of thyroid function (Table II). Immunization of BALB/c mice with TBP
Mice were immunized with either a soluble TBP or TBP-293 cells
(Fig. 1C). Native conformation of TBP was confirmed by its
ability to bind both TSH (Fig. 2) and patient autoantibodies (Table I).
Sera from immunized and control mice were tested against purified TBP
in an ELISA (Fig. 5A). Normal
mouse sera showed little or no binding (<1:100) to TBP, whereas, mice
immunized with either purified TBP or 293-TBP cells (groups 2 and 4,
respectively) showed considerable Ab response (1:12800). Similarly,
mice primed with either TBP or hETGP, and subsequently challenged with
293-TBP cells (groups 3 and 5, respectively) also showed significant Ab
response (>1:12800). Sera from these mice showed considerable TBII
activity (mean of 34.8–44.3%, relative to 3.84% for controls) (Fig. 5B and Table III). Moreover,
sera from affected mice induced enhanced levels of cAMP production in
CHO cells expressing human TSHR (Fig. 5C), with mean values
ranging from 16.1–24.2 pmol/ml for the test groups, compared with a
mean of 1.75 pmol/ml for controls (Table III). All mice in experimental
groups showed significant elevations in their T4 (Fig. 5D)
and T3 (Fig. 5E) levels. The mean T4 values for the
experimental groups ranged from 9.53 to 12.56 µg/dl, whereas the mean
for controls was 4.58 µg/dl (Table III). Mean T3 values ranged from
140.18–162.35 ng/dl, relative to a mean of 37.37 ng/dl in controls
(Table III). In addition, mice from experimental groups showed
considerably lower total body weight (Fig. 5F), with average
ranging from 20.6 to 21.67 g, whereas the controls weighed 24.2 g.
Together, these results showed that immunization of BALB/c mice with
the ectodomain of TSHR, either in a soluble form or expressed on
xenogeneic cells, can induce severe hyperthyroidism (Table III).
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As indicated in Fig. 6, a
majority of hyperthyroid mice, irrespective of TSHR preparation used
for immunization, showed enlargement of the thyroid gland relative to
thyroids from control mice due to hyperactivity (Fig. 6A).
Hematoxylin and eosin-stained sections from different regions of the
thyroid, harvested on day 120 after initiation of immunization, showed
hypertrophy and enlargement of colloids with thinning of the thyroid
epithelium (Fig. 6C). However, when thyroids collected from
affected mice on day 240 were examined for histopathology, some of them
showed focal necrosis and inflammation characterized by lymphocytic
infiltration (Fig. 6D). These observations are consistent
with features of Graves’ disease. Thyroids from control mice showed
normal histology (Fig. 6B).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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and Table II), there was no difference
in the ability of either TSHR protein to induce GD. The disease was
characterized by the appearance of Abs as detected in an ELISA followed
by the appearance of TBII and then TSAb activities. Subsequently, the
thyroid hormone levels were elevated. Consistent with these
observations, mice from most treatment groups showed a tendency toward
weight loss, with one group (group 4) showing a significant reduction
in weight. To our knowledge, this is the first report of induction of
Graves’ disease in nearly 100% of mice. Moreover, in the present
study, for the first time, the mouse TSHR protein was used to
successfully induce Graves’ disease in mice.
Our present study confirms and extends two earlier studies in which
RT4.15HP cells transfected with human TSHR were used for immunization
(13, 14). In our study, we were able to induce disease in
100% of mice, whereas in earlier studies (13, 14, 22, 23)
only 20% of the mice developed the disease. This, we believe, could be
due to several factors. In previous studies, RT4.15HP cells, which are
murine fibroblasts expressing MHC class-II and TSHR proteins, but
devoid of B-7 molecules, were used for immunization. On the other hand,
in the current study, M12 cells which are B cells that have been
extensively characterized as APCs and are known to express class-II and
B-7, were used (24). If M12 cells or RT4.15HP cells served
as APCs in vivo, then it is apparent that M12 cells would be more
efficient because, unlike RT4.15HP cells, they have the ability to
provide the second signal without which the immune response would be
suboptimal. Second, RT4.15HP cells had been transfected with a chimeric
class II construct containing 1, 2, and ß2 domains of the class
II molecule derived from H-2k, and ß1 domain
derived from H-2d (25). These cells
were originally constructed to identify functional domains of class II
molecule and had been used in vitro as APCs in T cell proliferation
assays to measure recall responses of T cells primed in vivo by direct
Ag inoculation (25). Expression of ß1 domain of
H-2d did not affect
H-2k-restricted APC function and suggested that
ß1 domain was not important for either peptide binding or for
recognition by T cells. RT4.15HP cells transfected with TSHR were used
to immunize AKR (H-2k) mice, and it is not clear
whether alloreactive responses against the chimeric class II molecule
rather than optimal TSHR-specific T cell responses were induced since
alloantigens are more potent immunogens than self Ags. Third, it is
likely that BALB/c (H-2d) mice used in this
study, relative to AKR mice used in the earlier studies (13, 14), might be more susceptible to GD. In fact, several earlier
studies have shown strain-specific differences in immune response to
TSHR (5, 23, 26). Fourth, it is well known that BALB/c
mice most often mount Th2 type of CD4+ T cell
responses (27) and that Th2 cells provide appropriate help
to B cells for high affinity Ab production. In this context, it is
interesting to note that CTB promotes activation of Th2 cells;
therefore, we used it as an adjuvant to see whether disease can be
induced in a higher proportion of mice. Since we were able to induce GD
in 100% of the mice without CTB (Table II, groups 6 and 7), the
effects of CTB, if any, could not be evaluated in this study. In fact,
Kita et al. (14) have reported that immunization of AKR
mice with RT4.15HP cells expressing TSHR along with the pertussis
toxin, a Th2 response-inducing adjuvant, caused disease in a higher
proportion (
50%) of animals. Since GD is an Ab-mediated disease, at
the present time it is not clear whether differential activation of
Th1- or Th2-type of CD4+ T cells in different
strains of mice contributed to differential susceptibilities.
Currently, efforts are underway in our laboratory to induce GD in
different strains of mice using CTB as an adjuvant.
Earlier, we and others have clearly shown that the ectodomain of TSHR is sufficient for TSH as well as stimulatory Ab binding (3, 4, 17, 19, 20, 22, 28, 29). Moreover, earlier studies showed that immunization of mice with fibroblasts expressing either TSHR or Class II alone was incapable of inducing an effective immune response against TSHR (13). However, upon immunization with cells expressing both TSHR and Class II molecules, mice developed autoantibody response to TSHR (13). This was taken to suggest that aberrant Class II expression might be critical for disease induction (13, 23). To test whether soluble ectodomain was sufficient to induce the disease and to address the requirement of TSHR-expressing cells to function as APCs, we conducted additional studies.
For these studies we used 293-TBP cells, from which soluble functional
TBP was purified. The 293 cells are human in origin and xenogeneic to
mice. As shown by our results (Fig. 5 and Table III), immunization of
BALB/c mice with either purified TBP or TBP-293 cells resulted in
development of severe hyperthyroidism in all mice except one. These
studies clearly demonstrated that soluble ectodomain of TSHR is
sufficient for disease induction in BALB/c mice. Previously, we used
ETSHR, hETGP, or mETGP produced in insect cells (3, 4, 10) to
immunize mice. These mice developed moderate hyperthyroidism but failed
to show classical symptoms often associated with human GD (5, 26, 30).
This, we believe, is due to lack of native conformation resulting from
aggregation of protein produced in insect cells. Although insect cell
lysates contain a very small amount of appropriately folded TSHR (as
demonstrated by neutralization of TBII activity in patient sera), most
of the protein remains aggregated. Since aggregated proteins are more
immunogenic than soluble proteins, a substantial Ab response was most
likely directed against epitopes on the aggregated protein, which may
or may not often be expressed on the native TSHR (30). In
contrast, TBP is readily soluble (16), and the purified
protein is fully functional in that it efficiently binds both TSH and
autoantibodies (16, 17) (Table I and Fig. 2). Current
studies further demonstrate that there is requirement neither for TSHR
to be cell associated nor for aberrant syngeneic Class II expression on
the cell surface. Since a typical APC requires expression of
costimulatory and adhesion molecules on its cell surface, it is not
apparent how TSHR-transfected RT4.15HP cells, which lacked
costimulatory molecules, could have served as APCs to prime
anti-TSHR responses. Moreover, RT4.15HP cells constitutively
expressed TSHR; therefore, they would be expected to process TSHR as an
endogenous Ag and present peptides bound to MHC class I. Since we know
that TSHR is a protein Ag, the immune response to it is likely to be
regulated by CD4+ T cells. These
CD4+ T cells often recognize antigenic peptides
bound to MHC class II, which requires exogenous Ag processing pathway.
Based on these observations and our current results, it is apparent
that TSHR ectodomain with native conformation is sufficient to cause GD
in 100% of immunized mice.
Now that we have a fully developed animal model for GD, further studies could lead to the development of new therapeutic strategies that could either down-modulate the immune response or shift it away from the production of pathogenic Abs.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Bellur S. Prabhakar, Professor and Head, Department of Microbiology and Immunology, College of Medicine, University of Illinois at Chicago, Room E 705, Medical Sciences Building, 835 South Wolcott Avenue, Chicago, IL 60612. E-mail address:
3 Abbreviations used in this paper: GD, Graves’ disease; TSHR, thyrotropin receptor; mTSHR, mouse TSHR; hTSHR, human TSHR; ETSHR, nonglycosylated ectodomain of hTSHR; mETGP, glycosylated ectodomain of mTSHR; hETGP, glycosylated ectodomain of hTSHR; mM12, M12 cells expressing mTSHR; hM12, M12 cells expressing hTSHR; CHO, Chinese hamster ovary cells; T4, tetraiodothyronine; T3, triiodothyronine; LTR, long terminal repeat; 293 cells, human embryonic kidney cells; bTSH, bovine thyrotropin; TBII, thyrotropin-binding inhibition index; CTB, cholera toxin B; TBP, soluble ectodomain of hTSHR produced in 293 cells.
Received for publication June 9, 1999. Accepted for publication August 19, 1999.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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promoter an efficient and versatile mammalian cDNA expression system composed of the simian virus 40 early promoter and the R-U5 segment of human T-cell leukemia virus type-I long terminal repeat. Mol. Cell. Biol. 8:466.
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 C. Vasu, B. S. Prabhakar, and M. J. Holterman Targeted CTLA-4 Engagement Induces CD4+CD25+CTLA-4high T Regulatory Cells with Target (Allo)antigen Specificity J. Immunol., August 15, 2004; 173(4): 2866 - 2876. [Abstract] [Full Text] [PDF]
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K. J. Land, J. S. Moll, M. H. Kaplan, and G. S. Seetharamaiah Signal Transducer and Activator of Transcription (Stat)-6-Dependent, But Not Stat4-Dependent, Immunity Is Required for the Development of Autoimmunity in Graves' Hyperthyroidism Endocrinology, August 1, 2004; 145(8): 3724 - 3730. [Abstract] [Full Text] [PDF]
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T. Muehlberg, J. A. Gilbert, P. V. Rao, A. M. McGregor, and J. P. Banga Dynamics of Thyroid-Stimulating and -Blocking Antibodies to the Thyrotropin Receptor in a Murine Model of Graves' Disease Endocrinology, April 1, 2004; 145(4): 1539 - 1545. [Abstract] [Full Text] [PDF]
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P. N. Pichurin, O. Pichurina, R. C. Marians, C.-R. Chen, Terry. F. Davies, B. Rapoport, and S. M. McLachlan Thyrotropin Receptor Knockout Mice: Studies on Immunological Tolerance to a Major Thyroid Autoantigen Endocrinology, March 1, 2004; 145(3): 1294 - 1301. [Abstract] [Full Text] [PDF]
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C.-R. Chen, P. Pichurin, G. D. Chazenbalk, H. Aliesky, Y. Nagayama, S. M. McLachlan, and B. Rapoport Low-Dose Immunization with Adenovirus Expressing the Thyroid-Stimulating Hormone Receptor A-Subunit Deviates the Antibody Response toward That of Autoantibodies in Human Graves' Disease Endocrinology, January 1, 2004; 145(1): 228 - 233. [Abstract] [Full Text] [PDF]
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B. S. Prabhakar, R. S. Bahn, and T. J. Smith Current Perspective on the Pathogenesis of Graves' Disease and Ophthalmopathy Endocr. Rev., December 1, 2003; 24(6): 802 - 835. [Abstract] [Full Text] [PDF]
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L. Schwarz-Lauer, P. N. Pichurin, C.-R. Chen, Y. Nagayama, C. Paras, J. C. Morris, B. Rapoport, and S. M. McLachlan The Cysteine-Rich Amino Terminus of the Thyrotropin Receptor Is the Immunodominant Linear Antibody Epitope in Mice Immunized Using Naked Deoxyribonucleic Acid or Adenovirus Vectors Endocrinology, May 1, 2003; 144(5): 1718 - 1725. [Abstract] [Full Text] [PDF]
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 C. Vasu, S. R. Gorla, B. S. Prabhakar, and M. J. Holterman Targeted engagement of CTLA-4 prevents autoimmune thyroiditis Int. Immunol., May 1, 2003; 15(5): 641 - 654. [Abstract] [Full Text] [PDF]
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R.-N. E. Dogan, C. Vasu, M. J. Holterman, and B. S. Prabhakar Absence of IL-4, and Not Suppression of the Th2 Response, Prevents Development of Experimental Autoimmune Graves' Disease J. Immunol., February 15, 2003; 170(4): 2195 - 2204. [Abstract] [Full Text] [PDF]
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T. Ando, M. Imaizumi, P. Graves, P. Unger, and T. F. Davies Induction of Thyroid-Stimulating Hormone Receptor Autoimmunity in Hamsters Endocrinology, February 1, 2003; 144(2): 671 - 680. [Abstract] [Full Text] [PDF]
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P. V. Rao, P. F. Watson, A. P. Weetman, G. Carayanniotis, and J. P. Banga Contrasting Activities of Thyrotropin Receptor Antibodies in Experimental Models of Graves' Disease Induced by Injection of Transfected Fibroblasts or Deoxyribonucleic Acid Vaccination Endocrinology, January 1, 2003; 144(1): 260 - 266. [Abstract] [Full Text] [PDF]
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 G. Mace, M. Jaume, C. Blanpied, L. Stephan, J. D. Coudert, P. Druet, and G. Dietrich Anti-{micro}-opioid-receptor IgG antibodies are commonly present in serum from healthy blood donors: evidence for a role in apoptotic immune cell death Blood, October 16, 2002; 100(9): 3261 - 3268. [Abstract] [Full Text] [PDF]
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Y. Nagayama, M. Kita-Furuyama, T. Ando, K. Nakao, H. Mizuguchi, T. Hayakawa, K. Eguchi, and M. Niwa A Novel Murine Model of Graves' Hyperthyroidism with Intramuscular Injection of Adenovirus Expressing the Thyrotropin Receptor J. Immunol., March 15, 2002; 168(6): 2789 - 2794. [Abstract] [Full Text] [PDF]
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J. Pritchard, N. Horst, W. Cruikshank, and T. J. Smith Igs from Patients with Graves' Disease Induce the Expression of T Cell Chemoattractants in Their Fibroblasts J. Immunol., January 15, 2002; 168(2): 942 - 950. [Abstract] [Full Text] [PDF]
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 J. G. Cundiff, S. Kaithamana, G. S. Seetharamaiah, J. R. Baker Jr., and B. S. Prabhakar Studies Using Recombinant Fragments of Human TSH Receptor Reveal Apparent Diversity in the Binding Specificities of Antibodies That Block TSH Binding to Its Receptor or Stimulate Thyroid Hormone Production J. Clin. Endocrinol. Metab., September 1, 2001; 86(9): 4254 - 4260. [Abstract] [Full Text] [PDF]
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A. W. C. Kung, K. S. Lau, and L. D. Kohn Epitope Mapping of TSH Receptor-Blocking Antibodies in Graves' Disease That Appear during Pregnancy J. Clin. Endocrinol. Metab., August 1, 2001; 86(8): 3647 - 3653. [Abstract] [Full Text] [PDF]
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P. Pichurin, X.-M. Yan, L. Farilla, J. Guo, G. D. Chazenbalk, B. Rapoport, and S. M. McLachlan Naked TSH Receptor DNA Vaccination: A TH1 T Cell Response in Which Interferon-{gamma} Production, Rather than Antibody, Dominates the Immune Response in Mice Endocrinology, August 1, 2001; 142(8): 3530 - 3536. [Abstract] [Full Text] [PDF]
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J.-L. Fan, O. Memar, D. J. McCormick, and B. S. Prabhakar BALB/c Mice Produce Blister-Causing Antibodies Upon Immunization with a Recombinant Human Desmoglein 3 J. Immunol., December 1, 1999; 163(11): 6228 - 6235. [Abstract] [Full Text] [PDF]
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