| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
ARTICLES |
Departments of Medicine and Pharmacology (R.P.M.), Mount Sinai School of Medicine, New York, New York 10029
Address all correspondence and requests for reprints to: Dr. T. F. Davies, Box 1055, Mount Sinai Medical Center, 1 Gustave L. Levy Place, New York, New York 10029.
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
These results demonstrated the successful production of large quantities of well characterized, biologically active, mTSHR-ecd antigen. In addition, the data showed that although the ectodomain of the mTSHR bound TSH, intact holoreceptor may be required for high affinity ligand binding. Whether the transmembrane region is required for direct ligand binding, as seen for other G protein-linked receptors, or whether it is needed to stabilize the ligand binding to the ectodomain and maintain a correctly folded state, remains unclear.
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
In addition to serving as a bioassay for hTSHR-Abs, the mouse exhibits thyroid antigen-induced experimental autoimmune thyroiditis. This model has become one of the most common tools for understanding the mechanism of autoimmune dysfunction (8). In the past, both thyroglobulin and thyroid peroxidase have been shown to initiate experimental autoimmune thyroiditis, using either human and/or mouse antigens (9, 10, 11). Recently, attempts to initiate autoimmune thyroid disease with hTSHR-ecd antigen have been largely unsuccessful. While one group has claimed success (12), we and others have failed to initiate thyroiditis, although marked immune responses to the immunizing antigen have been obtained (13, 14, 15). Such data again suggested that the mouse TSHR may be immunologically distinct from the human TSHR.
To address these questions it is necessary to characterize the mTSHR and evaluate its immunological effects. We have succeeded in producing large quantities of eukaryotic mouse TSHR ectodomain and have characterized highly purified preparations for binding of TSH ligand and TSHR-Abs.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Amplification and sequencing of mTSHR-ecd
The mTSHR-ecd (bp 61–1245) was amplified by RT-PCR using CBA/J
murine thyroid cDNA as the template, Vent DNA polymerase (New England
Biolabs, Beverly, MA), and the following mTSHR-ecd-specific primers:
forward (5'-GATCGGATCCGGCAAAGAGTGTGCGTCTCC) and reverse
(5'-GATCGGTACC
GAATTCTTATCAGTGATGGTGATGGTGATGTCCACCTCCCCTGTA
GCCCATGATATCTTCAC). These primers included the underlined
BamHI and EcoRI restriction sites. The reverse
primer comprised the mTSHR-ecd-specific complementary sequence,
followed by a 9-bp spacer sequence (double underlined)
coding for three glycine residues, an 18-bp sequence (in
boldface) coding for six histidines (6Hi), and a synthetic
stop codon (in italics). The 6Hi at the C-terminus allowed
further purification of the mTSHR-ecd-6Hi using
nickel-nitrilo-triacetic (Ni-NTA) resin chromatography. After
amplification, the murine thyroid DNA was digested with
BamHI and EcoRI, electrophoresed on 1% agarose
gels, purified from excised gel slices using glassmilk (GeneClean kit,
BIO 101, La Jolla, CA), and ligated into
BamHI/EcoRI-digested eukaryotic pAcGP67B
expression vector (PharMingen, San Diego, CA) to generate the
pAcGP67B-mTSHR-ecd plasmid. After restriction analysis, the mTSHR-ecd
was sequenced using the dideoxynucleotide termination method (Sequenase
kit, U.S. Biochemical Corp., Cleveland, OH).
Expression of mTSHR-ecd in Hi-5 insect cells
mTSHR-ecd cDNA was inserted into the BamHI cloning
site of the pAcGP67B vector downstream to the polyhedrin promoter and
gp67 signal peptide (17). Recombinant pAcGP67B/mTSHR-ecd plasmid was
cotransfected with Baculo-Gold DNA into sf9 insect cells as previously
described (18). The resulting recombinant virus was then used to infect
Hi-5 silkworm cells (19). Expression of mTSHR-ecd protein was assessed
by subjecting unpurified total cell lysates, particulate-insoluble
fractions, and cytosolic (water-soluble) fractions obtained after
sonication to SDS-PAGE followed by Coomassie blue staining (20 µg
total protein/lane) and Western blot analysis (5–10 µg total
protein/lane), as previously described (20, 21). Blots were developed
using either murine monoclonal antibody (mAb A10, supplied by Dr. J. P.
Banga, University of London), recognizing hTSHR residues 22–35 (21),
or rabbit antiserum (R 397) generated against an extreme C-terminus
hTSHR peptide (residues 397–415) (22), both at a 1:10,000 dilution.
Bound antibodies were detected using enhanced chemiluminescence (ECL
kit, Amersham, Aylesbury, UK).
Purification of the mTSHR-ecd
Using Ni-NTA-resin chromatography (Quiagen, Chatsworth, CA), the
mTSHR-ecd was purified either from total cell lysates of infected
insect cells (under denaturing conditions followed by on-column
refolding) or from cytosolic fractions (under native conditions). For
purification under denaturing conditions followed by refolding, insect
cells harvested 65 h postinfection were solubilized in 6
M guanidine-HCl followed by centrifugation (12,000 rpm for
30 min at 15 C). For analytical purification, 8.5 mg total supernatant
protein were mixed with 200 µl 50% Ni-NTA resin. For larger scale
purification, 20–50 mg total protein lysate were mixed with 0.5–1 ml
50% Ni-NTA resin. After batch binding for 1 h at room
temperature, the slurry was loaded onto low pressure chromatography
columns (Bio-Rad, Richmond, CA) and allowed to settle, and the resin
was sequentially washed with 10 vol 6 M guanidine-HCl (pH
8.0), 10 vol 8 M urea (pH = 8.0), and 40 vol 8
M urea (pH 6.5). The mTSHR-ecd captured by the Ni-NTA resin
was further subjected to refolding in situ, by applying two
consecutive linear gradients of refolding buffers (50–75 ml each),
over 10–12 h at 4 C: gradient 1 consisted of buffer A [8
M urea, 10 mM Tris-HCl (pH 7.4), 0.5
M NaCl, 20% glycerol, and 0.1% Triton X-100], and buffer
B was similar to buffer A, but contained 1 M urea. Gradient
2, started with buffer B and ended with buffer C [10 mM
Tris-HCl (pH 7.4), 0.15 M NaCl, 0.1 M potassium
acetate, and 0.1% Triton X-100]. The refolded mTSHR-ecd was then
eluted with 10 bed volumes of 350 mM imidazole in refolding
buffer C. Samples from the input, flow-through, sequential washes, and
eluted mTSHR-ecd fractions were precipitated with trichloroacetic acid,
dissolved in SDS-PAGE sample buffer containing 2% ß-mercaptoethanol
(ßME), and fractionated by SDS-PAGE. Purification was evaluated
initially by Coomassie blue staining (5–10 µg protein/lane) and
immunoblotting with the murine mAb A10 and R 397 antiserum (1–4 µg
protein/lane). The purified mTSHR-ecd fractions were pooled; dialyzed
extensively against 10 mM Tris (pH 7.4), 0.05 M
NaCl, and 0.1% Triton X; and concentrated with polyethylene glycol
(Mr, 20,000) to 0.25–0.5 mg/ml.
For purification of soluble mTSHR-ecd under native conditions, insect cell pellets were resuspended in sonication buffer [50 mM sodium phosphate (pH 7.4), 0.3 M NaCl, and protease inhibitor cocktail (PharMingen, San Diego, CA; catalog no. 21426Z)], and after two freeze-thaw cycles, the pellet was extracted on ice with three sonication cycles (1 min each) at 15-min intervals. Subsequently, the lysate was centrifuged at 50,000 x g for 40 min, and the soluble (cytosolic) fraction was filtered through a 0.2-µm filter. Filtrate (6–10 mg) was mixed with 250–500 µl Ni-NTA resin in a binding buffer containing 0.5 M NaCl, 10 mM ßME, 20% glycerol, and 60 mM imidazole, followed by batch binding for 2 h at 4 C. After casting the column, the resin was extensively washed with binding buffer without ßME. Bound cytosolic mTSHR-ecd was then eluted with 300 mM imidazole. Purity assessment and dialysis of pooled fractions were performed as described above.
Deglycosylation of purified mTSHR-ecd
Both refolded and cytosolic mTSHR-ecd were subjected to
deglycosylation with endoglycosidase F (Endo F) or endoglycosidase H
(Endo H; Boehringer Mannheim, Indianapolis, IN). mTSHR-ecd samples were
reduced by boiling for 3 min in the presence of 0.2% SDS, 1% ßME
for Endo F and 0.05% SDS-2% ßME for Endo H, and aliquots (5 µg
protein) were diluted into 10 vol Endo F buffer [20 mM
potassium phosphate (pH 7.5), 10 mM EDTA, and 1% Nonidet
P-40]) or Endo H buffer [50 mM sodium phosphate (pH 5.5)
and 0.5% Nonidet P-40]. After enzyme addition the samples were
incubated for 16 h at 37 C. Proteins were precipitated with 10%
trichloroacetic acid, rinsed with acetone, and dissolved in SDS-PAGE
sample buffer containing 2% ßME. The samples were subjected to
SDS-PAGE (1 µg/lane) followed by immunoblotting as described
above.
Binding of TSH to purified mTSHR-ecd
The ability of the mTSHR-ecd to bind TSH was tested indirectly
using a competition assay. In the first step, increasing amounts of
affinity-purified mTSHR-ecd protein (competitor) and
[125I]TSH (8,000 cpm; gift from Kronus, San Clemente, CA)
were combined in 300 µl binding buffer (NaCl-free Hanks’ Balanced
Salt Solution and 280 mM sucrose) for 1 h at 23 C.
Secondly, this mixture was added to JP-09 Chinese hamster ovary (CHO)
cells stably expressing the wild-type human TSHR (23) (gift from Dr. G.
Vassart, Free University of Brussels, Brussels, Belgium), and
incubation was continued for 1.5 h at 37 C, as previously
described (24). To explore the influence of folding of the mTSHR-ecd on
TSH binding, in separate experiments the refolded mTSHR-ecd was
reduced, unfolded by incubation with 10 mM dithiothreitol
followed by alkylation of the sulfhydryl groups of cysteine with 40
mM iodoacetamide, as described previously (25), and
incubated as described above. After incubation, the cells were washed
three times with ice-cold binding buffer dissolved in 0.5 ml 1
N NaOH, and the bound radioactivity was counted.
Solubilized porcine thyroid membranes enriched for TSHR (gift from
Kronus) served as a positive receptor control and was used at total
protein concentrations of 1.2–120 µg (estimated to contain 0.06–6
ng pTSHR). Wild-type baculovirus-infected insect cell lysate, used at
similar protein concentrations as the mTSHR-ecd, served as the negative
control. Competition by unlabeled bovine TSH (bTSH; Thytropar, Armour,
Kankakee, IL), used at 10-6-10-12
M, validated the assay. To confirm binding of ligand to the
recombinant receptor and not binding of receptor to CHO cells, the
refolded mTSHR-ecd was first preincubated with the JP-09 cells, and
[125I]TSH was added after washing the cells. Throughout,
bovine TSH was assumed to have a specific activity of 30 IU/mg.
Binding of hTSHR-Ab to recombinant mTSHR-ecd
Enzyme-linked immunosorbent assays were performed as previously
described (15). Briefly, 96-well microtitration plates (Immulon 2,
Dynatech Laboratories, Chantilly, VA) were coated with 100 µl/well
purified refolded mTSHR-ecd (1 µg/ml) or partially purified insect
cell-expressed human TSHR-ecd (18), refolded by gradual dilution of
denaturing agents and dialysis as described previously (26), and used
at 10 µg/ml. Negative controls were wild-type baculovirus-infected
insect cell lysate and BSA (used at 1–10 µg/ml). Samples were
diluted in carbonate bicarbonate buffer (pH 9.6). After coating
overnight at 4 C, wells were washed, blocked, and incubated with an
internationally validated hTSHR-Ab-containing serum [MRC long acting
thyroid stimulator activity (LATS) Research Standard B, National
Institute for Biological Standards and Control, London, UK]
reconstituted to yield an activity of 7.5 mU/ml, corresponding to a
1:50 serum dilution. Sera from 14 additional patients with hyperthyroid
Graves’ disease, containing high levels of hTSHR-Ab as defined by
porcine RRA (27), and 6 normal sera (all at 1:50 dilutions) were also
tested individually for binding to the purified refolded mTSHR-ecd and
control antigens. In a separate experiment, the hTSHR-Ab positive LATS
standard serum was also reacted in 2-fold serial dilutions, against
refolded and unfolded mTSHR-ecd. Three well characterized murine
monoclonal antibodies to the hTSHR-ecd (mAb A10, recognizing amino
acids 22–35; mAb A9 recognizing amino acids 147–229; and mAb A7,
recognizing amino acids 402–415) (21) were used as positive controls
at a 1:50,000 dilution. Detection was accomplished using alkaline
phosphatase-labeled antihuman IgG (A-5403, Sigma Chemical Co., St.
Louis, MO) and antimouse IgG (A-0532, Sigma Life Sciences), and
absorption was measured after 1 h at 405 nm.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
, the mTSHR-ecd cDNA probe
hybridized to four mTSHR mRNAs with estimated lengths of 4.3, 3.5, 2.9,
and 1 kilobases (kb; lane 2) present in the thyroid mRNA of euthyroid
CBA/J mice (group A). In this experiment, the 2.9-kb transcript was the
most abundant. The mouse cDNA probe also hybridized to three hTSHR
mRNAs in a human thyroid sample (lane 1); these migrated at 4.3, 1.7,
and 1.3 kb and were previously characterized by our laboratory (28). In
this sample, the 4.3-kb transcript was the most abundant. A second
Northern blot of murine thyroid mRNA, extracted from CBA/J mice treated
with methimazole and a low iodine diet to induce TSH stimulation (group
B), demonstrated a similar set of four mTSHR transcripts, also with the
2.9-kb transcript the most abundant (Fig. 1B, lane 1). No mTSHR-ecd
transcripts were detected in mRNAs extracted from mouse brain, muscle,
or liver (Fig. 1B, lanes 2–4).
|
) had a high substitution probability among proteins of
common ancestry, as evaluated using a matrix of relative substitution
frequencies (29). This frequency of substitution could have been due to
simple polymorphisms and/or mouse strain (BALB/c vs. CBA/J)
differences. Compared to the sequence of the hTSHR-ecd, 62 amino acid
differences were noted (Table 1).
|
) and R 397 (Fig. 2B), infection of Hi-5 insect cells with recombinant baculoviruses
containing mTSHR-ecd cDNA induced high level expression of two major
mTSHR-ecd proteins with apparent molecular masses of 66 kDa
(glycosylated) and 52 kDa (corresponding to the predicted molecular
size of unglycosylated mTSHR-ecd). Both species were detected in the
total cell lysate (Fig. 2, A and B, lane 1), the particulate-insoluble
fraction (Fig. 2, A and B, lane 2), and the cytosolic (water-soluble)
fraction (Fig. 2, A and B, lane 3). As evaluated by Coomassie blue
staining, these proteins were enriched in the insoluble fraction (not
shown). Very small amounts were detectable by Western blot in the
culture medium despite the presence of the baculovirus gp67 signal
peptide on the nascent mTSHR-ecd proteins (not shown). Nevertheless,
significant amounts (>10%) of the mTSHR-ecd species were present in
the cytosolic fraction. This enabled us to purify natively folded
mTSHR-ecd present in this fraction and compare it with mTSHR-ecd
purified from total cell lysates, followed by in vitro
refolding. Neither of the detected species was present in similar cell
fractions obtained from wild-type baculovirus-infected insect cells
(not shown).
|
, A and C), the mTSHR-ecd was purified
to homogeneity after Ni-NTA column elution, being most enriched in
eluted fractions 3 and 4 (lanes 5 and 6). The purified mTSHR-ecd again
comprised two major species, migrating at 66 and 52 kDa, and lower
amounts of partially glycosylated forms, migrating between 52–66 kDa
(Fig. 3B). Each of the mTSHR-ecd species identified on Coomassie blue
staining was recognized by mAb A10 in Western blots. The facts that mAb
A10 is recognizing an epitope at the extreme NH2-terminus (amino acids
22–35), and the C-terminus was immobilized by the Ni-NTA resin before
elution confirm the identity and integrity of all mTSHR-ecd species
purified. The total cell lysate (starting material), flow-through, and
last high stringency wash are shown in lanes 1–3 of Fig. 3, A and
C.
|
Purification of the cytosolic mTSHR-ecd. A similar analysis of
soluble mTSHR-ecd, purified using native conditions, is shown in Fig. 4. As shown by Coomassie blue staining (Fig. 4A), the
purified cytosolic mTSHR-ecd migrated as a unique broad band around 66
kDa and was enriched in the eluted fractions 3 and 4 (lanes 2 and 3). A
control represented by an enriched unpurified insoluble fraction from
insect cells expressing the mTSHR-ecd is shown in lane 4. This fraction
contained the 66-kDa mTSHR-ecd as a major protein and also lower mol wt
contaminant proteins, absent in lanes 2 and 3 containing the purified
mTSHR-ecd. Immunoblotting with mAb A10 confirmed the Coomassie blue
data by identifying the purified mTSHR-ecd fractions (lanes 4–7). The
unpurified cytosolic fraction (input), flow-through, and last wash are
shown in lanes 1–3. Under native conditions, the 66-kDa major
glycoform was predominant in the purified fractions of cytosolic
mTSHR-ecd compared with the purification under denaturing/refolding
conditions, where a spectrum of lower mol wt glycosylated
intermediates and the 52-kDa nonglycosylated form were also purified
(Fig. 4B, lane 8). Using this method, 100-µg amounts of cytosolic
mTSHR-ecd could be purified, representing 0.2–0.5% of the total
protein input.
|
, lanes 1 and 2). The smaller
amounts of partially glycosylated intermediates (not as evident here)
were also reduced. Similar results were obtained after digestion with
Endo H, which is known to cleave only high mannose sugars (Fig. 5A, lanes 3 and 4). Figure 5B shows that these enzymes had the same effect
on the 66-kDa glycosylated species purified from the cytosol using
native conditions. These deglycosylation patterns demonstrate that no
significant differences could be found in the glycosylation status of
the two mTSHR-ecd preparations, and that most of the carbohydrate
side-chains were of the high mannose type. This finding is not
unexpected, as it is known that insect cells are able to add high
mannose oligosaccharides to N-acetylglucosamine residues
linked to asparagine and to partially trim these structures to shorter
oligosaccharides, but are generally not able to add complex sugars such
as galactose and sialic acid to the trimmed sugar backbone (19).
|
103 mIU/L; Fig. 6B, inset). Solubilized porcine TSHR, known
to bind TSH with high affinity (28), also inhibited the binding of
[125I]TSH to the hTSHR expressed in CHO cells in a
dose-dependent manner (Fig. 6A), confirming the validity of the assay.
Based on quantitative data derived from Western blot analyses, nanogram
amounts of the porcine TSHR were effective at inhibiting
[125I]TSH binding, achieving 50% inhibition at a
concentration of 5 x 10-11 M. Both
the purified cytosolic and refolded mTSHR-ecd similarly inhibited
labeled TSH binding in a dose-dependent manner. The inhibition induced
by the two preparations had almost identical affinities, confirming the
functional activity of both preparations (Fig. 6A). Although
concentrations of 2 x 10-7 M for the
cytosolic and 3.3 x 10-7 M for the
refolded receptor produced almost complete inhibition, 50% inhibition
was achieved by concentrations of 4–5 x 10-8
M (Fig. 6A). Compared to the inhibitory effect of unlabeled
bTSH, the affinity of the purified mTSHR-ecd was lower by about 1 log.
The wild-type baculovirus-infected insect cell protein lysate
(subjected to similar mock procedures as the purified mTSHR-ecd) did
not compete for [125I]TSH binding to the hTSHR-CHO cells
(Fig. 6A). When the refolded mTSHR-ecd was preincubated with the
hTSHR-CHO cells and then removed before the addition of
[125I]TSH, no binding inhibition was achieved, confirming
the direct binding of the recombinant mTSHR-ecd preparations to
[125I]TSH. In contrast to results with the cytosolic and
refolded mTSHR-ecd species, denatured and unfolded mTSHR-ecd failed to
compete for the binding of [125I]TSH at similar protein
concentrations (Fig. 6A). The unfolded mTSHR-ecd preparation was intact
and migrated at the correct molecular mass, as evaluated by
immunoblotting with mAb A10 and mAb A7 (not shown). These data
demonstrated the critical importance of the folding state of the
mTSHR-ecd for TSH binding.
|
and 8), mAb A7
(amino acids 402–415; not shown), and mAb A10 (amino acids 22–35;
Fig. 7B). The functional relevance of the refolded mTSHR-ecd
immunoreactivity was further demonstrated by the marked degree of
binding of a standard Graves’ TSHR-Ab-positive serum (expressed in MRC
LATS Standard B units) and tested in 1:50 and 1:100 serum equivalent
dilutions (Fig. 7A). This standard serum also recognized an hTSHR-ecd
preparation, but did not react with wild-type baculovirus-infected
insect cell control protein lysate or BSA (Fig. 7A). The recognition of
recombinant mTSHR-ecd by hTSHR-Ab was confirmed by the examination of
14 additional serum samples from patients with hyperthyroid Graves’
disease with high titers of hTSHR-Ab (as determined by competition with
labeled TSH for binding to porcine TSHR). The purified refolded
mTSHR-ecd interacted with 4 of the 14 Graves’ sera (28%), but did not
react with any of 6 normal control sera negative for hTSHR-Ab (Fig. 8).
None of the Graves’ or normal sera reacted with control antigens (Fig. 8).
|
|
). In contrast,
reduction of the folded ectodomain markedly diminished, but did not
abolish, the binding of standard Graves’ TSHR-Ab (Fig. 7C).
Approximately 75% of the activity was lost when expressed in relation
to mAb A10 binding, indicating that the folding state of the mTSHR-ecd
was an important determinant of the degree of hTSHR-Ab binding. |
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Using a baculovirus-insect cell system, we synthesized and purified the murine TSHR-ecd and characterized the product as a test antigen. Compared to our previous data (18) and those of other workers (32, 33), we obtained a much higher degree of purity as well as quantity of soluble recombinant TSHR-ecd. Affinity-purified antigen was present as two major species [glycosylated (66 kDa) and unglycosylated (52 kDa)] as well as minor, partially glycosylated forms. The purified mTSHR-ecd inhibited binding of radiolabeled TSH to CHO cells expressing functional hTSHRs and provided indirect evidence of TSH binding to the recombinant product. The fact that the refolded mTSHR-ecd inhibited [125I]TSH binding with a similar affinity as the natively folded (cytosolic) mTSHR-ecd confirmed the correct refolding of this preparation. However, the level of inhibition was less than that attained using solubilized porcine TSHR. This lower reactivity could be partly explained by the lower affinity of TSH for the mTSHR (our unpublished observations). Indeed, native mTSHR has been shown to exhibit 1 order of magnitude lower affinity for TSH compared to the hTSHR (5).
There are at least two alternative explanations for the lower affinity
of the mTSHR-ecd for TSH; the absence of the transmembrane region and
differences in the glycosylation status of the recombinant product.
Although it is generally agreed that the large ectodomain of
glycoprotein receptors is primarily involved in ligand binding, and the
transmembrane domain is involved in signal transduction, the issue of
whether these domains must interact for high affinity binding remains
open. Recent studies have shown that the hyt/hyt mutation in
the mTSHR (Pro556 Leu in the fourth transmembrane region),
abolished TSH binding and receptor function (34). Similarly, mutation
of Asp383 to Asn in the second transmembrane region of the
LH receptor (35), significantly lowered the affinity of lutropin ligand
for its receptor. These studies suggest that the transmembrane region
may be important for direct high affinity ligand binding or may
indirectly stabilize the ligand-ectodomain binding complex. However,
studies comparing binding of TSH by the 
-subunit of the hTSHR
(lacking the transmembrane region) and solubilized full-length hTSHR of
transfected cells (36) could not find significant ligand binding
affinity differences, although these binding affinities were
inexplicably low (10-5 M). Similarly, studies
comparing binding of LH to insect cell-expressed full-length and
truncated LH receptors (37) could not detect significant differences,
although both were 1 order of magnitude less than the natural ovarian
LH receptor. Further studies are clearly necessary to clarify the role
of the transmembrane region in ligand binding. As far as the
glycosylation status of the mTSHR-ecd was concerned, most of the insect
cell-expressed murine ectodomain was glycosylated in a high mannose
state. It has previously been demonstrated that TSH was able to bind,
albeit with a lower affinity, to insect cell-expressed,
nonglycosylated, refolded hTSHR-ecd (26). In contrast, Zhang et
al. (37) have shown that tunicamycin-treated insect cells
expressing a carbohydrate-free LH receptor lacked ligand-binding
activity, as opposed to insect cell LH receptor glycosylated in a high
mannose state. Hence, the LH and TSH receptors may differ in their
requirements for ligand binding. Our data showed that the high mannose
glycosylation of the mTSHR-ecd did not hinder TSH binding. Indeed,
differential enzymatic cleavage of insect cell-expressed LH receptor
has shown that only the proximal N-acetylglucosamine
residue, common to both high mannose and complex carbohydrate
side-chains, is necessary for acquisition of a correctly folded
receptor that binds LH with high affinity (37). Therefore, the folding
status seems to be more important than glycosylation in ligand
binding.
We thought that for physiological characterization, a soluble, correctly folded recombinant mTSHR-ecd product was necessary. Obtaining a soluble form of recombinant TSHR-ecd has been problematic. Harsh preparative techniques were originally used to render prokaryotic products soluble (20), although new procedures for hydrophilic products have been recently described (38). With eukaryotic insect cell systems, however, either no soluble product was found or only very small quantities of soluble receptor were identified after extensive purification (32). In the present studies, 100-µg amounts of soluble cytosolic mTSHR-ecd were obtained, enough to allow a quantitative, direct comparison to the initially insoluble forms that were rendered soluble by in vitro refolding. Both the molecular sizes and degrees of glycosylation appeared identical, and both products competed with radiolabeled TSH with equal avidity. The unfolding of the mTSHR-ecd resulted in marked diminution of TSH binding to the recombinant product. Hence, the final folding status of the product was more important than the initial solubility status. The purified mTSHR-ecd also bound some, but not all (30%), of the human TSHR autoantibodies present in the sera of selected patients with hyperthyroid Graves’ disease. In contrast with the monoclonal antibodies, human autoantibodies from patients with Graves’ disease had a higher affinity for the folded state, indicating the presence of antibodies to important conformationally dependent epitopes. Similar results were obtained from structure-activity studies of thyroid peroxidase autoantigen and thyroid peroxidase antibodies (25). Unfolding the mTSHR-ecd did not totally abolish autoantibody recognition, as 25% of the binding remained. Hence, linear epitopes are also viable interaction sites for human TSHR-Abs, in agreement with their significant, but low, affinity recognition of prokaryotic antigen (18, 20).
In conclusion, we have succeeded in generating large quantities of biologically active mTSHR-ecd. However, compared to the natural TSHR, the binding of TSH was of lower affinity and was recognized by only 30% of hTSHR-Abs from patients with Graves’ disease. The fact that only a minority of TSHR autoantibodies bound both natural TSHR and recombinant mTSHR-ecd suggested that structural differences existed between them. Such differences may have been secondary to the lack of the transmembrane region in the recombinant product, variation in the glycosylation status, or differences in the protein structures of the ectodomains. These structural dissimilarities are likely to generate fine changes in nonlinear epitopes, which may only be mapped knowing the crystallographic structure of the molecule.
|
Acknowledgments |
|---|
|
Footnotes |
|---|
2 Present address: University of Nagasaki, Nagasaki, Japan.
3 Theodore and Florence Baumritter professor.
Received October 21, 1996.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
This article has been cited by other articles:
 |
|
 |
|
  S. M. McLachlan, Y. Nagayama, and B. Rapoport Insight into Graves' Hyperthyroidism from Animal Models Endocr. Rev., October 1, 2005; 26(6): 800 - 832. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 Y. Oda, J. Sanders, S. Roberts, M. Maruyama, A. Kiddie, J. Furmaniak, and B. Rees Smith Analysis of Carbohydrate Residues on Recombinant Human Thyrotropin Receptor J. Clin. Endocrinol. Metab., June 1, 1999; 84(6): 2119 - 2125. [Abstract] [Full Text]
|
|
|
|
 |
|
 M. Kita, L. Ahmad, R. C. Marians, H. Vlase, P. Unger, P. N. Graves, and T. F. Davies Regulation and Transfer of a Murine Model of Thyrotropin Receptor Antibody Mediated Graves' Disease Endocrinology, March 1, 1999; 140(3): 1392 - 1398. [Abstract] [Full Text]
|
|
|
|
 |
|
 Y. Nagayama, H. Namba, N. Yokoyama, S. Yamashita, and M. Niwa Role of Asparagine-linked Oligosaccharides in Protein Folding, Membrane Targeting, and Thyrotropin and Autoantibody Binding of the Human Thyrotropin Receptor J. Biol. Chem., December 11, 1998; 273(50): 33423 - 33428. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B. Rapoport, G. D. Chazenbalk, J. C. Jaume, and S. M. McLachlan The Thyrotropin (TSH)-Releasing Hormone Receptor: Interaction with TSH and Autoantibodies Endocr. Rev., December 1, 1998; 19(6): 673 - 716. [Abstract] [Full Text]
|
|
|
|
|
|
C. R. Da Costa and A. P. Johnstone Production of the Thyrotrophin Receptor Extracellular Domain as a Glycosylphosphatidylinositol-anchored Membrane Protein and Its Interaction with Thyrotrophin and Autoantibodies J. Biol. Chem., May 8, 1998; 273(19): 11874 - 11880. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Osuga, S.-G. Liang, J. S. Dallas, C. Wang, and A. J. W. Hsueh Soluble Ecto-Domain Mutant of Thyrotropin (TSH) Receptor Incapable of Binding TSH Neutralizes the Action of Thyroid-Stimulating Antibodies from Graves' Patients Endocrinology, February 1, 1998; 139(2): 671 - 676. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. R. Baker Jr Autoimmune Endocrine Disease JAMA, December 10, 1997; 278(22): 1931 - 1937. [Abstract] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Endocrinology | Endocrine Reviews | J. Clin. End. & Metab. |
| Molecular Endocrinology | Recent Prog. Horm. Res. | All Endocrine Journals |
