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Thyroid Division Department of Medicine Brigham and Women’s Hospital Harvard Medical School Boston Massachusetts 02115
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIAL AND METHODSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIAL AND METHODSREFERENCES |
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In vertebrate cells, basal and substrate-induced selective enzyme proteolysis is also involved in the control of transformation of T4 to the active hormone T3 , the first step in thyroid hormone action. Two enzymes, the types 1 (D1) and 2 (D2) iodothyronine deiodinases, can catalyze this reaction. Along with type 3 deiodinase (D3) that inactivates T4 and T3, these integral membrane selenoproteins constitute a homeostatic system that controls the intracellular concentration of active thyroid hormone within human tissues. Whereas D1, expressed primarily in liver and kidney, is considered to be the major source of circulating T3, D2 catalyzes the local production of T3 in the central nervous system, pituitary gland, and brown adipose tissue (5). As a result, thyroid hormone receptor (TR) occupancy by T3 is higher in these tissues than is the case in cells in which plasma is the only source of intracellular T3. In addition, the effects of a decrease in plasma T4, such as occurs in iodine deficiency, are mitigated by a rapid compensatory increase in D2 activity (5). One key feature of D2 that allows such plasticity is its short half-life (<1 h) (6).
Considerable evidence indicates that D2 regulation by its substrates is posttranslational (7, 8, 9) and recent data implicate the proteasome system (10, 11). Indeed, in pituitary tumor cells, proteasomal inhibitors (MG132 or lactacystin) stabilize D2 activity for several hours in the presence of cycloheximide (CX) or the D2 substrates T4 or rT3 (10). Parallel reductions of transiently expressed 75Se-labeled D2 and D2 activity occur after exposure of cells to CX and/or rT3, indicating that this reduction is due to catabolism of the protein rather than to an alteration in its structure (11). Single amino acid changes in the active center of D2, which either raise the Michaelis-Menten constant (Km) for substrate approximately 1,000-fold or block its catalytic activity, impair or eliminate substrate-induced loss of D2 (11). These results suggest that interaction of substrate with D2 selectively targets the protein for degradation by the proteasome system. Despite this, an ubiquitinated D2 intermediate has not been identified nor has the role of catalysis in its production been clarified.
Proteins containing selenocysteine (Sec) are synthesized slowly due to the complex mechanisms required to suppress the stop codon function of the Sec codon, UGA (12). This and its short half-life result in low cellular D2 concentrations. D2 protein contains few immunogenic peptide sequences, resulting in difficulties in the generation of high-affinity antibodies suitable for Western blots (11). Accordingly, to enhance the possibility of detecting ubiquitinated D2, we employed a mutated D2 enzyme in which Cys is substituted for Sec 133 in the active center. This protein is transiently expressed at levels approximately 100-fold higher than the native D2 but is catalytically active and subject to the same substrate regulatory pathways as is the native enzyme (11). We also labeled D2 with FLAG, an epitope for which there is a highly specific antibody. By transiently expressing catalytically active FLAG-tagged D2, we have identified ubiquitinated-D2 and show that its levels are increased by exposure of cells to substrate. We also found that the Ub-activating enzyme, E1, is required for Ub conjugation to D2 and for the substrate-induced acceleration of that process. Surprisingly, while we have confirmed previous results showing that substrate also reduces D1 activity, this T4 activating enzyme is not ubiquitinated. This indicates that ubiquitination of D2 is a specific mechanism that confers rapid posttranslational regulation by T4 of its activation to T3. This is a unique example of substrate-induced selective proteolysis that involves ubiquitination of an ER resident protein. To our knowledge, it is the first demonstration that such a regulatory pathway controls activation of a hormone.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIAL AND METHODSREFERENCES |
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). In three independent experiments the
D2 activity increased an average of approximately 30% at the
restrictive temperature (Fig. 1B), whereas no changes were detected in
similarly treated wild-type CHO cells (data not shown).
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). These treatments caused an
approximately 20% reduction in D2 activity in cells at 30 C
(P < 0.05). However, at 40 C, these agents had no
significant effect on D2 activity (P > 0.05
vs. time 0; P < 0.05 vs. 30 C)
(Fig. 1, D–E). The effects of CH or rT3 in
wild-type CHO cells transiently expressing D2 were not affected by
incubation at 40 C (data not shown). These results indicate that E1 is
rate limiting in the basal and substrate-induced changes in D2
activity.
To confirm that there was a correlation between enzyme activity
and D2 protein we evaluated changes of
75Se-labeled D2 under the same conditions. Paired
plates of ts20 cells transiently expressing wild D2 were labeled with
Na2[75Se]O3,
and 24 h later one plate was shifted to 40 C for 4 h (Fig. 2A). The plate lysates were processed for
immunoprecipitation (IP) using D2 antiserum and the precipitates
analyzed by SDS-PAGE. The 31-kDa 75Se-D2 band
increased 1.8 ± 0.6 fold (P < 0.05) in cells at
40 C vs. those at 30 C, again consistent with impaired
ubiquitination (and subsequent proteolysis) of D2 at the restrictive
temperature (Fig. 2, B and C) as the explanation for the increased D2
activity. However, no 75Se-labeled higher
molecular mass proteins were visualized in the control samples as
would be expected if ubiquitination of D2 was occurring under these
circumstances (Fig. 2B).
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) were treated with 100
µM CX or 30 µM rT3,
and sonicates were processed for Western blotting with anti-FLAG
antibody (Fig. 4A). The 
32 kDa
FLAG-NH2-cysD2 band behaved similarly to
the 31 kDa 75Se-D2 band detected in the IP
of HEK-293 cells reported previously (11). It disappeared with a
half-life of approximately 2 h in the presence of CX and was also
decreased by treatment with rT3. Both effects
were blocked by concomitant exposure to 10 µM MG132, an
inhibitor of proteasomal proteolysis (Fig. 4B). D2 activity paralleled
the level of FLAG-NH2-cysD2 protein, as
visualized by the Western blot (Fig. 4, B and C). Most importantly,
five high molecular mass bands (100–300 kDa) were observed in the
Western blot, consistent with the formation of polyubiquitinated D2
(Fig. 4D). These bands constituted 10–20% of the immunoreactive FLAG
protein in the cells.
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). Three 110- to 300-kDa bands,
similar in size to those seen with the anti-FLAG antibody in Fig. 4D, reacted with the anti-Ub antiserum. Stripping the anti-Ub antibodies
from the blot and reprobing it with anti-FLAG antibody confirmed that
these proteins were ubiquitinated FLAG-cysD2 (Fig. 4E). Microsomal and
cytosolic fractions were prepared from sonicates of HEK-293 cells, and
FLAG-cysD2 as well as Ub-FLAG-cysD2 were found to be localized
predominantly in the particulate fraction (Fig. 4F).
If the high molecular mass FLAG-D2 proteins are ubiquitinated, then it
should be possible to block their formation by inactivating E1 in the
ts20 cells. Increasing the incubation temperature to 40 C for
2 h markedly reduced the amount of the high molecular mass
FLAG-NH2-cysD2 complexes in ts20 cells (Fig. 5A). There was also an associated
approximately 20% increase in 32 kDa FLAG protein and D2 activity
(100 ± 13 vs. 126 ± 18%; P <
0.05) in the same cells.
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), establishing that
substrate-enzyme interaction accelerates the rate of D2 ubiquitination.
As expected, the FLAG-NH2-cysD2 band decreased by
40–50% as did D2 activity (100 ± 20 vs. 45 ±
12%; P < 0.05). In a complementary experiment,
addition of 10 µM MG132 not only increased the
32 kDa FLAG-NH2-cysD2 band by 50–60% but
also increased the high molecular mass
FLAG-NH2-cysD2 bands, as would be predicted if
MG132 blocked their proteasomal uptake (Fig. 5C). The D2 activity of
these cells also increased parallel to the
FLAG-NH2-cysD2 bands (100 ± 17
vs. 155 ± 25%; P < 0.05).
The COOH Terminus Is Critical for the Proteolysis of Ub-D2
According to the current topological model for the deiodinases,
the NH2 terminus of D2 is located in the lumen of
the ER with its catalytic portion in the cytosol. As mentioned
previously, fusion of the FLAG sequence to the
NH2-terminus of D2 does not change its
degradation rate (Fig. 4B). Surprisingly, the levels of D2 activity
(Fig. 6A) and protein (Fig. 6B) were
4-fold higher in cells transiently expressing cysD2 with the FLAG
epitope placed at the COOH terminus (Fig. 3B). The amount of
Ub-FLAG-COOH-cysD2 was massively increased, about 20- to 30-fold,
suggesting that the degradation of Ub-D2 was blocked by the presence of
the COOH-terminal FLAG epitope (Fig. 6B). Studies with CX confirmed
that the half-life of the FLAG-COOH-cysD2 protein was longer than that
of the wild-type D2 or the FLAG-NH2-cysD2 (Fig. 6, C and D). While CX exposure caused an approximately 50% decrease in
the FLAG-NH2-cysD2 protein over 2 h,
FLAG-COOH-cysD2 protein disappeared much more slowly, with a predicted
half-life of 4–5 h. Parallel effects occurred in D2 activity.
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Substrate Causes D1 Inactivation but Not Ubiquitination
As mentioned, the selenoenzyme D1 also catalyzes
T4 to T3 conversion
although the Km(T4) for
this enzyme (2 µM) is approximately 1000-fold higher
than is that for D2 (1–2 nM) (14). While D1 has sequence
and, presumably, structural similarities to D2, it differs from the
latter in having a relatively long half-life (>12 h; Fig. 7A). Accordingly, we prepared a
FLAG-NH2-cysD1 protein (Fig. 3C) to allow
comparative studies of the degradation pathways of D1 and D2. The
FLAG-NH2-cysD1 was again catalytically active,
and the half-life of D1 activity of transiently expressed FLAG-cysD1
was >12 h, 6 times longer than that of FLAG-cysD2 (Fig. 7A). Despite
much higher transient expression of D1, there was no detectable
Ub-FLAG-NH2-cysD1 conjugate in the HEK-293 cells
(Fig. 7B). However, exposure to rT3 for 24 h
caused a 70–80% reduction in D1 activity but no change in the level
of FLAG NH2-cysD1 protein (Fig. 7, C and D).
These results indicate that, despite the fact that substrate causes
decreases in both D2 and D1 activities, the mechanisms by which those
changes occur are quite distinct.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIAL AND METHODSREFERENCES |
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Two results suggest that ubiquitination and proteasomal degradation of
D2 initiate at the COOH terminus. First we can identify Ub-D2 in
particulate fraction of cell sonicates (Fig. 4F). The accumulation of
large amounts of Ub-D2 associated with the ER (20–50% of total FLAG
protein) indicates that the rate of D2 ubiquitination exceeds that of
Ub-D2 proteolysis. This agrees with the hypothesis that
proteolysis of ER proteins requires extraction of Ub-conjugated protein
from the ER (16). The alternative possibility that the COOH-terminal
portion of Ub-D2 is clipped off by an ER-bound proteasome, or by
regulated intramembrane proteolysis (18), is unlikely because small
FLAG-containing protein fragments (<32 kDa) were not detected (Fig. 6). Second, a comparison of the fate of NH2- and
COOH-FLAG-tagged D2 proteins allowed us to gain further insight into
the mechanistic aspects of D2 proteolysis. On one hand, conjugation of
FLAG to the NH2 terminus does not alter the
half-life of cysD2 (Fig. 4) (11). In contrast, the fusion of the FLAG
sequence to the COOH terminus of D2 not only prolonged its half-life
but also increased the size of the Ub-D2 pool 20- to 30-fold (Fig. 6).
Because both D2 activity and the 32 kDa protein were increased 3- to
4-fold in cells transfected with FLAG-COOH-cysD2, the increase in the
Ub-D2 pool is not due to increased ubiquitination. Rather, it is
probably caused by impaired proteasomal extraction/proteolysis of
Ub-D2. The accumulated Ub-D2 can then be recycled through Ub
isopeptidases to D2. Consequently, D2 half-life, protein levels, and
activity are all increased. This is supported by the data in Fig. 5.
Likewise, moving ts20 cells to the restrictive temperature decreases
Ub-D2 conjugates while it increases D2 protein and activity (Fig. 5).
The dynamic equilibrium between Ub-D2 and D2 also explains why D2
activity is stabilized for several hours by exposing CX-treated
pituitary tumor cells (10) or HEK-293 cells transiently expressing D2
(11) to MG132. The data presented in Fig. 5 also indicate that D2
activity parallels the levels of D2 protein and not Ub-D2 conjugates,
which, therefore, must be catalytically inactive. This indicates that
the D2 and Ub-D2 pools are in dynamic equilibrium that shifts toward D2
in the presence of MG132 or toward the formation of Ub-D2 when cells
are exposed to substrate. The immediate implication of these findings
is that proteasomal uptake can be a limiting step in D2 proteolysis, as
has been suggested for Sec62 (17).
Exposure to the substrate rT3 decreases D2
protein and activity by approximately 50%. The present results reveal
that this is due to an increase in D2 ubiquitination since both this
and the loss of D2 activity are blocked by E1 inactivation (Figs. 2 and 3). Because D2-substrate interaction is required to increase D2
proteolysis (11), it is possible that postcatalytic structural changes
in the D2 protein accelerate the ubiquitination cascade. Alternatively,
the redox state of the molecule might play a role. Reducing agents,
e.g. dithiothreitol, act as cofactors for D2 catalysis
in vitro by reducing the Se in the enzyme’s active center
after it is oxidized during the monodeiodination process (19).
Oxidation of the Se- in the native enzyme or SH
in the Cys mutant could be the primary signal that accelerates
ubiquitination. This would explain why an oxidizing agent such as
diamide irreversibly inactivates D2 (20). Indeed, such a
redox-sensitive mechanism has been described for the ubiquitination and
proteasomal degradation of hypoxia-inducible factor 1
, involved in
the activation of the erythropoietin gene (21). However, a
catalytically inactive D2 mutant, in which alanine was substituted for
the Sec in the active center of the enzyme (alaD2), retains the typical
D2 short half-life while it is refractory to the substrate-induced
acceleration of its proteolysis (11). If the redox state of the active
center plays a role in accelerating ubiquitination it must be limited
to the latter process.
The results with D1 provide an important contrast suggesting that
simply deiodinating an iodothyronine does not accelerate ubiquitination
of a deiodinase. The present results confirm previous data that D1
activity is also decreased by exposure to substrate (20), but our
results show that this does not involve D1 ubiquitination (Fig. 7).
Rather, the inactivation process is most likely the consequence of
oxidation of the active center Se that requires time until the
intracellular environment returns it to the reduced state, as
originally proposed (22). The finding that D1 is not ubiquitinated can
also explain its much longer half-life (Fig. 7). Taken together, these
results indicate that T4 to
T3 conversion by D2 is more tightly regulated
than that by D1. Whether this is due to differences in protein
structure or in subcellular localization between the two
selenodeiodinases remains to be determined.
In summary, we propose that newly synthesized D2
(D2-Se-) exists in the ER membrane in a dynamic
equilibrium with its ubiquitinated (inactive) derivative
Ub-D2-Se- (Fig. 8). The latter may be deubiquitinated by
isopeptidases, thus reactivating it, or it may enter proteasomes for
irreversible degradation. T4, the principal substrate of D2
[or rT3 (10, 11)], induces poorly understood changes
in the enzyme forming D2-Se, thus accelerating the ubiquitination
process, shifting the equilibrium toward Ub-D2-Se. Whether Ub-D2-Se can
be reactivated in vivo is not clear. The critical change
induced in D2-Se- by T4
constitutes a posttranslational control process regulating the rate of
T4 activation. Such a mechanism is especially
well suited to T4 since its 7-day plasma
half-life in humans precludes minute-to-minute variations due to
changes in secretion via feedback at the hypothalamic-pituitary level.
However, given the advantages of local intracellular control of hormone
activation, it would not be surprising if other hormone systems are
regulated in a similar fashion.
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MATERIAL AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIAL AND METHODSREFERENCES |
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Preparation of D1- and D2-Expressing Plasmids and
Mutagenesis
All constructs were cloned into the D10 mammalian expression
vector (23). Wild-type D2 constructs contained the SelP SECIS element
(24). Overlap-extension PCR was used to produce D2 mutants where the
Sec 133 was replaced by Cys (cys-D2), as described previously (11), and
subcloned into the same vector. A D1 mutant in which Cys was replaced
by Sec (G5; cysD1) was described previously (12).
Epitope-tagged D2 proteins were created using the eight-amino acid FLAG
sequence (Sigma). The NH2-FLAG-cysD2
(Met-Asp-Tyr-Lys-Asp-Asp-Asp-Asp-Lys Leu-Ala-Met) was
generated by the Bp84-Bp85 oligonucleotides [the former contained an
NcoI site, while the latter contained a SalI site
(lower case)] by Vent PCR on a hD2 Cys template. (Bp84
sense; 5'-catgccATGG GCATCCTCAG CGTAGACTTG CTGA; Bp85 antisense:
5'-ttccgcggcc gctatggccg acgtcgacTT AACCAGCTAA TCTAGTTTTC TTTCATCT).
The resulting fragment was cut by NcoI/SalI and
cloned into the corresponding sites of the D10 vector, which also
contained a 5'-EcoRI site and a strong Kozak consensus (-3A)
sequence with an in-frame NcoI site at the 3'-end of the
FLAG sequence (Fig. 3A). This construct does not contain a SECIS
element so that translation of the NH2-FLAG-cysD2
terminates at the second UGA codon, the eighth codon from the COOH end
of the native D2 protein. Previous studies showed no effect of this
truncation on enzyme activity (25). The COOH-FLAG-cysD2 Cys mutant was
generated by the Bp97-Bp95 oligonucleotides [the former contained an
EcoRI site, while the latter contained an XbaI
site (lower case)] by Vent PCR on a hD2 Cys template. (Bp97
sense: 5'-ggaattcatt ATGGGCATCC TCAGCGTAGA CTTGCTGATC A; Bp95
antisense: GCTCTAGAtt acttgtcgtc atcgtccttg tagtcACCAG CTAATCTAGT
TTTCTTaCAT CTCTTGCT). The sense oligo contained the same Kozak
consensus as the NH2-FLAG-cysD2 construct, to ensure the
same rate of translational initiation. We also replaced the second Sec
codon (UGA) with Cys (UGU) using the Bp95 antisense oligo, to ensure
uniform translation (Fig. 3B). Both FLAG-cysD2 constructs were
catalytically active.
The NH2-FLAG-cysD1 mutant was generated by the Bp92-Bp93
hD1 oligonucleotides on a rat Cys mutant template (NOREF>G5), incorporating
NheI and SalI restriction sites (lower
case) at the 5'- and 3'-ends of cysD2, respectively (Bp92 sense;
ctagctagcc ATGGGGCTGC CCCAGCCAGG GCTGTGGCTG A; Bp93 antisense:
ttccgcggcc gctatggccg acgtcgacTT AACTGTGGAG CTTTTCCAGA ACAGCACGA). The
fragment was cut by these enzymes and cloned between the corresponding
sites of a version of the above described NH2-FLAG D10
fusion vector, containing a unique NheI site 3' to the FLAG
sequence (Fig. 3C). The resulting protein is a rat
NH2-FLAGcysD1 containing Pro, Pro, and Gly in positions 4,
6, and 7, respectively, as in the human wild-type D1. Its COOH terminus
contains His and Ser in positions 248–249 and is followed by a stop
codon, making it a 29-kDa protein. The protein was catalytically
active. None of the FLAG-tagged Cys mutant deiodinase fusion proteins
contained a SECIS element. The accuracy of the construction was
confirmed by sequencing.
Procedures for Transfections
D1 and D2 were transiently expressed by introducing expression
vectors containing the respective cDNAs into human embryonic kidney
epithelial cells (HEK-293) or Chinese hamster ovary (CHO) ts20 cells.
To obtain uniform expression in all plates in an experiment, we used
the following pair-plating approach to the transfection. Cells were
initially plated in 60-mm dishes and grown until confluence in DMEM
supplemented with 10% FBS. Plasmid DNA was precipitated in ethanol and
then redissolved in 0.25 M CaCl2 in
HEPES buffer and equal amounts of suspension added to the pair of
plates. Ten micrograms of D10 vector containing D1 or D2 cDNAs were
combined with 4 µg of a D15 vector and 3 µg of TKGH plasmid per
plate. Cells and plasmid DNA were allowed to stand for 20–30 min at
room temperature and subsequently incubated at 37 C. hGH was measured
in the media 48 h later as a monitor for transfection efficiency.
Differences in hGH expression in cells transfected with the same DNA
precipitate were <5%.
75Se incorporation and D2 IP
These procedures were performed as previously described (11).
Briefly, transfected ts20 cells were labeled in vivo with
4–6 uCi of
Na2[75Se]O3/dish
on day 2 after transfection in the presence of DMEM supplemented with
10% FBS. On day 3, the cells were lysed for 2–3 h at 4 C using a
lysis buffer [1% Triton X100, 1% bovine hemoglobin, 1
mM iodoacetamide, 0.2 U aprotinin/ml, 1
mM phenylmethylsulfonyl fluoride (PMSF) in TSA
buffer (0.01 M Tris-HCl, pH 8.0, 0.14
M NaCl, 0.025% NaN3)] 1
ml/dish. After centrifugation of the lysate at 1,000 rpm for 5 min,
each 1 ml supernatant was incubated for 12–24 h at 4 C with preimmune
rabbit sera to a final dilution of 1:100. One hundred microliters of a
10% Pansorbin suspension were then added per tube and incubated under
slow agitation for 20 min at 4 C. After centrifugation at 1,000 x
g for 7 min, the supernatants were incubated for 24–48 h at
4 C with a D2 rabbit polyclonal antibody [No. 85254 (11)] to a final
dilution of 1:100. This antibody was generated against a synthetic
peptide SRSKSTRGEWRRMLTSEGLRC (residues 52–72) selected from the human
D2 protein. Immunoprecipitates were obtained after the addition of 100
µl of a 10% Pansorbin suspension and centrifugation at 1,000 rpm for
7 min. The pellets were then washed four times with a dilution buffer
(0.1% Triton X 100, 0.1% bovine hemoglobin in TSA) and then washed
once in TSA buffer and once with 0.05 M Tris-HCl,
pH 6.8. Pellets were then heated at 95 C for 7 min in sample loading
buffer and spun at top speed for 5 min, and 30 µl of the supernatants
were used for analysis by SDS-PAGE.
Western Blots of Epitope-Tagged D2
HEK-293 or ts20 cells transiently expressing the various
constructs were scraped, washed in PBS, suspended in lysis buffer (0.01
M Tris-HCl, pH 8.0, 0.14 M NaCl, 0.25
M sucrose, 1 mM PMSF, 2 µg/ml aprotinin, 2
µg/ml leupeptin), and sonicated for 3–4 sec. In one experiment cell
sonicates were centrifuged at 2,500 x g for 10 min,
and the supernatant was spun at 100,000 g for 1 h to
separate microsomes from cytosol. Protein concentration was measured as
described by Bradford (26), and 20–40 µg were analyzed by 7.5% or
12% SDS-PAGE and electrotransferred to a polyvinylidene fluoride
membrane (Immobilon, Millipore Corp.,
Bedford, MA). The blots were probed with an anti-FLAG M2 antibody
(1:3333, Sigma) or with a polyclonal anti-Ub antibody
(1:1000, Chemicon, Temecula CA), or both. In the latter case, Ub
detection was followed by stripping and exposure to the same FLAG
detection. Samples processed for the Ub antibody were previously
purified on M2-anti-FLAG affinity agarose (Sigma). The
Western blot was carried out using the Chemiluminescence Kit of
Roche Molecular Biochemicals (Indianapolis IN), according
the instructions of the manufacturer.
Statistical Analysis
Data are presented as mean ± SD
throughout the studies. Student’s t test was used for
comparative analysis. Five percent was the level of significance
required to reject the null hypothesis.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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This work was supported by NIH Grant RO1-DK-36256. A.C.B. was partially supported by the University of Sao Paulo (Sao Paulo, Brazil).
Received for publication June 15, 2000. Revision received August 8, 2000. Accepted for publication August 10, 2000.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIAL AND METHODSREFERENCES |
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 W. S. da-Silva, J. W. Harney, B. W. Kim, J. Li, S. D.C. Bianco, A. Crescenzi, M. A. Christoffolete, S. A. Huang, and A. C. Bianco The Small Polyphenolic Molecule Kaempferol Increases Cellular Energy Expenditure and Thyroid Hormone Activation Diabetes, March 1, 2007; 56(3): 767 - 776. [Abstract] [Full Text] [PDF]
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F. Goulart da Silva, G. Giannocco, M. F. Santos, and M. T. Nunes Thyroid Hormone Induction of Actin Polymerization in Somatotrophs of Hypothyroid Rats: Potential Repercussions in Growth Hormone Synthesis and Secretion Endocrinology, December 1, 2006; 147(12): 5777 - 5785. [Abstract] [Full Text] [PDF]
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 A. Zeold, L. Pormuller, M. Dentice, J. W. Harney, C. Curcio-Morelli, S. M. Tente, A. C. Bianco, and B. Gereben Metabolic Instability of Type 2 Deiodinase Is Transferable To Stable Proteins Independently of Subcellular Localization J. Biol. Chem., October 20, 2006; 281(42): 31538 - 31543. [Abstract] [Full Text] [PDF]
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S. Yasuo, N. Nakao, S. Ohkura, M. Iigo, S. Hagiwara, A. Goto, H. Ando, T. Yamamura, M. Watanabe, T. Watanabe, et al. Long-Day Suppressed Expression of Type 2 Deiodinase Gene in the Mediobasal Hypothalamus of the Saanen Goat, a Short-Day Breeder: Implication for Seasonal Window of Thyroid Hormone Action on Reproductive Neuroendocrine Axis Endocrinology, January 1, 2006; 147(1): 432 - 440. [Abstract] [Full Text] [PDF]
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C. H. A. Gouveia, M. A. Christoffolete, C. R. Zaitune, J. M. Dora, J. W. Harney, A. L. Maia, and A. C. Bianco Type 2 Iodothyronine Selenodeiodinase Is Expressed throughout the Mouse Skeleton and in the MC3T3-E1 Mouse Osteoblastic Cell Line during Differentiation Endocrinology, January 1, 2005; 146(1): 195 - 200. [Abstract] [Full Text] [PDF]
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S. Yasuzawa-Amano, N. Toyoda, A. Maeda, A. Kosaki, Y. Mori, T. Iwasaka, and M. Nishikawa Expression and Regulation of Type 2 Iodothyronine Deiodinase in Rat Aorta Media Endocrinology, December 1, 2004; 145(12): 5638 - 5645. [Abstract] [Full Text] [PDF]
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F. W. J. S. Wassen, W. Klootwijk, E. Kaptein, D. J. Duncker, T. J. Visser, and G. G. J. M. Kuiper Characteristics and Thyroid State-Dependent Regulation of Iodothyronine Deiodinases in Pigs Endocrinology, September 1, 2004; 145(9): 4251 - 4263. [Abstract] [Full Text] [PDF]
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 A. L. Fellet, P. Arza, N. Arreche, C. Arranz, and A. M. Balaszczuk Nitric oxide and thyroid gland: modulation of cardiovascular function in autonomic-blocked anaesthetized rats Exp Physiol, May 1, 2004; 89(3): 303 - 312. [Abstract] [Full Text] [PDF]
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 B. W. Kim, A. M. Zavacki, C. Curcio-Morelli, M. Dentice, J. W. Harney, P. R. Larsen, and A. C. Bianco Endoplasmic Reticulum-Associated Degradation of the Human Type 2 Iodothyronine Deiodinase (D2) is Mediated via an Association between Mammalian UBC7 and the Carboxyl Region of D2 Mol. Endocrinol., December 1, 2003; 17(12): 2603 - 2612. [Abstract] [Full Text] [PDF]
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 O. Broedel, M. Eravci, S. Fuxius, T. Smolarz, A. Jeitner, H. Grau, G. Stoltenburg-Didinger, H. Plueckhan, H. Meinhold, and A. Baumgartner Effects of hyper- and hypothyroidism on thyroid hormone concentrations in regions of the rat brain Am J Physiol Endocrinol Metab, September 1, 2003; 285(3): E470 - E480. [Abstract] [Full Text] [PDF]
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S. Friedrichsen, S. Christ, H. Heuer, M. K. H. Schafer, A. Mansouri, K. Bauer, and T. J. Visser Regulation of Iodothyronine Deiodinases in the Pax8-/- Mouse Model of Congenital Hypothyroidism Endocrinology, March 1, 2003; 144(3): 777 - 784. [Abstract] [Full Text] [PDF]
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C. Curcio-Morelli, B. Gereben, A. M. Zavacki, B. W. Kim, S. Huang, J. W. Harney, P. R. Larsen, and A. C. Bianco In Vivo Dimerization of Types 1, 2, and 3 Iodothyronine Selenodeiodinases Endocrinology, March 1, 2003; 144(3): 937 - 946. [Abstract] [Full Text] [PDF]
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M. Baqui, D. Botero, B. Gereben, C. Curcio, J. W. Harney, D. Salvatore, K. Sorimachi, P. R. Larsen, and A. C. Bianco Human Type 3 Iodothyronine Selenodeiodinase Is Located in the Plasma Membrane and Undergoes Rapid Internalization to Endosomes J. Biol. Chem., January 3, 2003; 278(2): 1206 - 1211. [Abstract] [Full Text] [PDF]
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D. Botero, B. Gereben, C. Goncalves, L. A. de Jesus, J. W. Harney, and A. C. Bianco Ubc6p and Ubc7p Are Required for Normal and Substrate-Induced Endoplasmic Reticulum-Associated Degradation of the Human Selenoprotein Type 2 Iodothyronine Monodeiodinase Mol. Endocrinol., September 1, 2002; 16(9): 1999 - 2007. [Abstract] [Full Text] [PDF]
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B. Gereben, A. Kollar, J. W. Harney, and P. R. Larsen The mRNA Structure Has Potent Regulatory Effects on Type 2 Iodothyronine Deiodinase Expression Mol. Endocrinol., July 1, 2002; 16(7): 1667 - 1679. [Abstract] [Full Text] [PDF]
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A. C. Bianco, D. Salvatore, B. Gereben, M. J. Berry, and P. R. Larsen Biochemistry, Cellular and Molecular Biology, and Physiological Roles of the Iodothyronine Selenodeiodinases Endocr. Rev., February 1, 2002; 23(1): 38 - 89. [Abstract] [Full Text] [PDF]
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R. Peeters, C. Fekete, C. Goncalves, G. Legradi, H. M. Tu, J. W. Harney, A. C. Bianco, R. M. Lechan, and P. R. Larsen Regional physiological adaptation of the central nervous system deiodinases to iodine deficiency Am J Physiol Endocrinol Metab, July 1, 2001; 281(1): E54 - E61. [Abstract] [Full Text] [PDF]
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C. Curcio, M. M. A. Baqui, D. Salvatore, B. H. Rihn, S. Mohr, J. W. Harney, P. R. Larsen, and A. C. Bianco The Human Type 2 Iodothyronine Deiodinase Is a Selenoprotein Highly Expressed in a Mesothelioma Cell Line J. Biol. Chem., August 3, 2001; 276(32): 30183 - 30187. [Abstract] [Full Text] [PDF]
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