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Thyroid Division, Department of Medicine, Brigham and Women’s Hospital and Harvard Medical School, Boston, Massachusetts 02115
Address all correspondence and requests for reprints to: P. Reed Larsen, M.D., Professor of Medicine, Thyroid Division, Department of Medicine, Brigham and Women’s Hospital and Harvard Medical School, Boston, Massachusetts 02115. E-mail: Larsen{at}rascal.med.harvard.edu
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Abstract |
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Introduction |
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Type 2 deiodinase activity was first identified in pituitary tissue from hypothyroid rats where it catalyzed the rapid production of significant quantities of specifically bound nuclear T3 following i.v. infusion of T4 by a PTU-insensitive reaction (6). Thyroxine causes a rapid suppression of TSH release from the thyrotroph, a process that is blocked by prior treatment of rats with iopanoic acid, a competitive inhibitor of D2 (7). While initially D2 was differentiated from D1 activity by its insensitivity to PTU (8, 9, 10), subsequent studies identified other major kinetic differences between D2 and D1. The apparent Km of D2 for T4 is approximately three orders of magnitude lower than that of D1 and D2 activity is increased, but D1 decreased, in hypothyroid animals (3, 5). The D2 messenger RNA (mRNA) and its activity in BAT is markedly increased by the adrenergic stimulation during exposure of rats to cold (5). Type 2 deiodinase is an obligate source of T3 for the rat central nervous system and pituitary, where it contributes over 50% of the specifically bound nuclear T3 (11). It thus can serve as the transducing mechanism by which the hypothalamic-pituitary axis can recognize the level of the prohormone T4 in the circulation. Recent in situ hybridization studies have provided the surprising result that D2 mRNA expression in the central nervous system is highly localized in the tanycytes lining the third ventricle, a result consistent with earlier activity studies that identified the arcuate nucleus median eminence portion of the rat hypothalamus as that portion of the CNS containing the highest concentration of D2 activity (12, 13, 14). This suggests that cerebrospinal fluid T4 could serve an important role in the feedback regulation of TSH synthesis and release by T3.
D2 activity is controlled at two levels in the pituitary and central nervous system. The best documented effect is that substrates for this enzyme, such as T4 and 3,3'5'-T3 (reverse T3) accelerate the degradation rate of the protein (15). Both T4 and reverse T3 are more potent in producing this effect than is T3 pointing to a nonnuclear mechanism (16, 17). In addition, D2 mRNA has recently been shown to be inversely proportional to thyroid status in the pituitary and the central nervous system (5, 12, 18).
We have recently developed a semiquantitative competitive RT-PCR technique for the quantitation of D2 and cyclophilin (Cyc) mRNAs, which permits the identification of the transcripts in small samples of tissue and applied this to the study of D2 mRNA regulation in the rat anterior pituitary and medial basal hypothalamus (12). The purpose of the present studies was to validate the RT-PCR technique and use it to elucidate the mechanism for the acute effects of thyroid and other hormones on D2 mRNA expression.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Cell culture
GC and GH4C1 cells were cultured using
Ham’s F10 media containing 14 mM sodium bicarbonate, pH
7.4, and 2 mM L-glutamine supplemented with
10% FBS (Gibco, Grand Island, NY) until cells reached to about
70–80% confluence (19). For studies of T3 effects,
the culture media was removed from cells, the cells were rinsed with
PBS once, and Ham’s F10 media containing 10% charcoal-treated
(T3 and T4 depleted) FBS was added. Charcoal
was used to remove glucocorticoid and thyroid hormones from serum. The
media was changed after 1 day and the incubation continued for a second
day. T3 (100 nM) was diluted in Ham’s F10
media+10% charcoal-treated FBS and added to cells for the indicated
times. 8-Bromoadenosine 3', 5'-cyclic monophosphate (0.3
mM, 8 Br-cAMP), 3-isobutyl-1-methylxanthine (0.5
mM, IBMX), actinomycin D (2 µM),
TSH-releasing hormone (TRH) (1–1000 nM), dexamethasone
(Dex, 100 nM), and phorbol 12-myristate 13-acetate (100
nM-1 µM, TPA) were from Sigma Chemical Co. (St. Louis, MO).
5' Deiodinase assays
D2 assay was performed as described previously (4). Briefly,
mixtures of about 200 µg cell protein, 0.1 nM 3,3',5,
125I-5' tetraiodothyronine (T4) purified by
LH-20 chromatography, 2 nM unlabeled T4, 20
mM DTT, and 1 mM PTU in PE buffer in a final
volume of 300 ul were incubated for 60 or 120 min at 37 C. Free
125I was separated from unreacted substrate or
iodothyronine products by trichloroacetic acid precipitation after
addition of 200 µl of horse serum as described (20). Deiodination was
linear with both protein concentration and time and the quantity of
enzyme assayed was adjusted to consume <30% of substrate. All
reactions were in triplicate.
Northern blot analysis
RNA was isolated from anterior pituitary (AP), medial basal
hypothalamus (MBH), and GH4C1 cells using
TRIzol reagent (Life Technologies, Gaithersburg, MD).
Twenty micrograms of total RNA were electrophoresed through 1%
agarose/1.85% formaldehyde/2 µg ethidium bromide/1 x
MOPS gel at room temperature according to the procedure of
Lehrach et al. (21). RNA was transferred by capillary action
or by vacuum blotter with 10 x SSC according to the
manufacturer’s recommendations (Genescreen Plus, NEN Life Sciences,
Boston, MA) and completion of transfer was verified by the presence of
all of the 0.24–9.5 kb RNA molecular weight marker bands (BRL,
Gaithersburg, MD) as well as the 28S and 18S rRNA. RNA was fixed to the
nylon membrane by UV cross-linking with a Stratalinker
(Stratagene, La Jolla, CA).
Blots were hybridized for D2 mRNA under high stringency conditions (40% formamide/2 x Denhardt’s reagent/0.64 M NaCl/40 mM PO4, pH 5.6/0.8 mM EDTA, pH 7.4/7% dextran sulfate/1.6% SDS/80 µg/ml denatured sonicated thymus DNA) at 42 C overnight after prehybridization. D2 probed Genescreen Plus filters were washed sequentially twice with 2 x SSC/0.1% SDS at RT, once with 0.25 x SSC/0.1% SDS at RT, and once with 0.25 x SSC/0.1% SDS at 42 C. Blots were autoradiographed for 3–7 days with an intensifying screen at -80 C.
A cDNA probe was made by random priming of template DNA with
[
-32P]-dCTP. A 0.95-kb fragment (bp 421 to bp 1371,
EcoRI digest) of the rD2 partial cDNA clone 5–1 in
Bluescript SK vector (rD2 pBSSK, kindly provided by D. St. Germain and
V. A. Galton) containing small portions of the 5' and 3' flanking
regions and 788 nt of coding region (5) was gel purified for the D2
template. Quantitation of hybridization signals was done with a
scanning densitometer or phosphorimager with Molecular Dynamics, Inc. (Sunnyvale, CA) software version 3.1.
Semiquantitative competitive RT-PCR
Semiquantitative RT-PCR was performed as previously described
but with modifications in the quantitative aspects of the technique
(12). RT was performed with rD2-specific antisense primer
[nucleiotides (nt) 1381 to 1358] and rCyc-specific antisense primer
(nt 557 to 534) using total RNA (22). For the competitive PCR, the rD2
partial cDNA clone 5–1 in Bluescript SK vector (rD2 pBSSK) was used to
prepare a mutant D2. The region between two BsaA I
restriction sites (nt 688 to nt 843) within the coding region was
deleted and religated, producing a fragment that was 156 bp shorter
than the wild-type (Fig. 1A
). A mutant
Cyc plasmid was also created from wild-type Cyc plasmid by removing an
internal NcoI fragment (nt 339 to 464) (Fig. 1B). The RT
reaction was performed with 1 µg of total RNA for the 20 µl
reaction as described previously (12). Four microliters of RT product
was used for the PCR, which was performed separately for D2 and Cyc. 26
cycles were applied to all samples except for D2 in GC cells (30
cycles) annealing at a temperature of 62 C. The quantity of mutant D2
added for competitive PCR was 1.56–6.25 pg for GC cells and 12.5–25
pg for the other samples, whereas 12.5 pg of mutant Cyc was used for
all reactions. The PCR reaction included four microliters of RT product
and 40 µCi 32P-[alpha]-dCTP and was performed with Vent
polymerase in a 40 µl reaction volume. After 5 min denaturing at 95
C, 26 (or 30) cycles of PCR were performed with a denaturing 1' at 95
C, annealing 1'30'' at 62 C and amplification 3' at 76 C, followed by
10' extension. After separate amplification of both the D2 and Cyc, the
PCR products were run through a 4.5% polyacrylamide gel or when
indicated, on a denaturing gel (see Results). The gel was
dried and analyzed by phosphorimager.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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). PAGE
analysis of the combined PCR products of the D2 and Cyc RT reactions
showed 7 labeled bands (Fig. 2). Separate
electrophoresis of the products of the PCR of D2 wt and mut plasmids
showed two bands of the predicted sizes, 806 (wt) and 650 (mut) plus a
third larger band (upper band) (Fig. 2). As the concentration of the wt
D2 was reduced, the quantity of the wt and upper bands decreased
progressively and the mut D2 band increased. To confirm that these
bands were wt and mut D2 amplicons, the 806 and 650 bp bands were
extracted from the gel and digested with BsaA I or BsaH I.
With BsaA I, three fragments (507, 156, and 143 bp) were produced
as predicted (Fig. 3). Digestion of mut
D2 by BsaA I produced two fragments of 507 and 143 bp again as
predicted (Figs. 1 and 3, lanes 2 and 5). Digestion of wt D2 by
BsaH I produced the predicted 562 and 244 bp fragments, but the
mut D2 was not restricted again as predicted. This result indicated the
authenticity of the D2 amplicons. The upper band was not restricted
with either enzyme, indicating it was likely to be a heteroduplex form
as has been reported by others (23).
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and 3). As with the
D2 PCR, the quantity of both wt and the upper bands decreased with
decreasing input wt cDNA and the mut Cyc band increased.
To determine the composition of the upper bands, they were extracted
from the gel, denatured, and subjected to PAGE under denaturing
conditions together with the wt and mut bands as markers. The upper
band from the D2 amplification resolved into equal amounts of two bands
corresponding to the sizes of the specifically amplified wt and mut D2
PCR products (Fig. 4, lanes 1–3).
Similar results were obtained with the upper bands from the Cyc
amplification (Fig. 4, lanes 4–6). This indicated that the upper bands
consisted of heteroduplexes of equal quantities of wt and mut strands.
The presence of the wild-type amplicon for both D2 and Cyc explained
the parallel decrease in the upper band and wt amplicons. Because the
upper bands consisted of equal parts of the wild-type and mutant bands,
50% of the density of the upper band must be added to that of wt and
mut bands to obtain an accurate assessment of the quantity of PCR
fragments formed during the reaction, although this ignores minor
differences in the C content of the wild-type and mutant bands.
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, 1–3) were performed for D2 and
Cyc using this pool as template. Subsequently, quantitative PCR was
performed in duplicate (A and B) using these separate three RT reaction
products (Fig. 5A). The bands produced from the 3 RT reactions gave the
same pattern as the standard (Fig. 5A). Standard curves were plotted
using the ratios of the corrected wt and mut amplicons (Fig. 5B). The
mean ±SD of D2/Cyc mRNA ratios from these six reactions
was 1.94 ± 0.139 (coefficient of variation of 7.2%), indicating
that the method is highly reproducible (Fig. 5B).
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). GH4C1 cells were exposed to 100
nM TRH for 24 h, with D2 mRNA expression increasing
about 30% according to both analyses (Table 2B). It should be noted that the D2/Cyc
ratios are 10-fold higher by the PCR than by Northern due to the more
efficient amplification of the D2 than of the Cyc cDNA. In all cases in
the studies described below, there was no alteration in Cyc mRNA during
any of the perturbations. Therefore, the changes in the D2/Cyc ratio
are a consequence of changes in D2 mRNA.
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). To analyze the time course and mechanism
for this effect, T3-responsive rat pituitary tumor cell
lines were evaluated. According to the competitive RT-PCR, GC cells
(D2/Cyc ratio of 0.42 ± 0.035, n = 3) cultured in 10% FBS
expressed about 10-fold less D2 mRNA than GH4C1 cells (4.62 ±
0.09, n = 3). In agreement with this, the D2 activity in GH4C1
cells (14 ± 0.58 fmol/minMG protein: mean ±
SE) was 8.4-fold higher than in GC cells (1.66 ±
0.13: mean ± SE). By Northern blotting, we observed a
strong signal in mRNA from GH4C1 cells, whereas we could not detect a
specific band in GC cell mRNA under the same conditions (data not
shown). To examine the effects of T3 on D2 mRNA, cells were
transferred to thyroid hormone and glucocorticoid depleted-FBS
[charcoal-stripped, (CS), FBS] for several days. This caused an
increase of 2.0- to 3.5-fold in the D2/Cyc mRNA ratios in both GH4C1
and GC cells over 48 h with the increase in GC cells being
consistently higher (about 3.3-fold). Accordingly, we used
"hypothyroid" GC cells to study the acute effects of T3
on D2 mRNA expression. Addition of T3 to hypothyroid GC
cells caused a rapid decrease in the D2/Cyc ratio which fell to 50% of
its initial value at 2 h (Fig. 6),
indicating that thyroid hormone deficiency accounts for the increase in
D2 mRNA during incubation in the hormone-depleted sera. To assess
whether the T3 effect was due to reduced transcription or
to an acceleration of D2 mRNA degradation, we analyzed the effects of
actinomycin D (Act D), with or without T3, on these
changes. Act D caused a rapid decrease in D2 mRNA (but no changes in
Cyc) with the D2/Cyc ratio again decreasing 50% within 2 h (Fig. 6A). When the treatments were extended to 4 h, T3 did
not decrease the D2/Cyc ratio further, whereas D2 expression continued
to decrease with Act D (Fig. 6B). T3 did not alter the
effect of Act D. Thus, T3 rapidly reduced D2 gene
transcription, but there is also a residual fraction of D2 mRNA
expression in GC cells which is independent of thyroid status.
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). Pretreatment with CHX for 30 min
followed by T3 for 2 h decreased D2 mRNA expression to
a similar degree, indicating that new protein synthesis is not required
for a T3-induced reduction in D2 mRNA transcription.
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). In contrast, 50 or
100 nM reverse T3 caused either a slight
increase (A) or no change in the D2/Cyc mRNA ratio (B), but an 80–90%
decrease in D2 activity (A). Thus, T3 decreases D2 activity
largely by suppression of transcription while the effect of reverse
T3 is exclusively posttranscriptional. Control studies
demonstrated that the effects of reverse T3 on D2 activity
were time dependent and thus not explained by dilution of the
T4 substrate with reverse T3 carried over into
the D2 activity assay.
Dexamethasone transiently increases D2 gene transcription
In contrast to the effect of T3, 100 nM
dexamethasone (Dex) increased D2 mRNA expression 2.5-fold in
hypothyroid GC cells but D2 mRNA decreased to near control levels after
2 h (Fig. 8). This transient increase of
D2 mRNA expression was completely blocked when GC cells were treated
with 100 nM Dex and 2 µM ActD together,
showing that the effect was transcriptional. When GC cells were treated
with 100 nM T3 and 100 nM Dex
together, the Dex effect was substantially reduced (Fig. 8).
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). GH4C1 cells were
also sensitive to 8 Br-cAMP/IBMX, although the effect was smaller than
in GC cells (Fig. 9A). This suggested that D2 mRNA expression was
modestly up-regulated by the cAMP signaling pathway in pituitary tumor
cells.
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).
TPA did not increase D2 mRNA expression. Exposure to TRH or TPA for
24 h caused a larger increase in D2 activity, which for 100
nM TRH was 77% and for TPA was 62%. |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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and 5). For D2, there is a single extra band and with
cyclophilin a doublet is found (Fig. 2). These bands are heteroduplex
combinations of mutant and wild-type cDNAs as identified in this and
earlier studies (Fig. 4) (23, 29). They are not susceptible to
restriction by the predicted enzymes as are the authentic wild-type and
mutant amplicons (Figs. 3 and 4). Analysis by denaturing PAGE indicates
that these complexes consist of equal quantities of mutant and
wild-type cDNA strands (Fig. 4). Because these are bona fide
products of the amplification reaction, they must be included in
determining the ratio of wild-type to mutant amplicons. This can be
achieved either by adding half of the density of the upper band to that
of the authentic wild-type and mutant cDNAs, or the PCR reaction
products can be denatured and analyzed by denaturing PAGE. We used the
former approach in these studies.
In competitive PCR approaches, a sample of mutant target mRNA can be
included in the initial RT reaction to allow correction for the
efficiency of the amplification reaction. This is required if absolute
quantitation, rather than a determination of the ratio of a variable
target mRNA (D2) to an unchanging internal control mRNA (Cyc) is
desired. Figure 5B shows that the RT reaction primed by gene-specific
D2 and Cyc oligonucleotides is reproducible such that the D2/Cyc mRNA
ratio in three different reactions from the same mRNA pool gives a
coefficient of variation of <10% (Fig. 5B). Because the various
short-term hormonal perturbations used in these studies do not change
the level of cyc mRNA, we used this approach. The excellent agreement
between changes in the D2/Cyc mRNA ratios by Northern analyses and
competitive PCR in anterior pituitary, medial basal hypothalamus, and
GH4C1 cells confirms the accuracy of the technique (Table 1). The
absolute ratio of D2/Cyc by Northern analysis is approximately 10-fold
lower than that obtained by RT-PCR. This indicates that the PCR
conditions result in 10-fold greater relative amplification of the D2
than of the Cyc cDNA. While there could be a number of reasons for this
related to the sequences or the quantities of oligonucleotides present,
it does not affect the analysis of changes in the D2/Cyc mRNA
ratios.
The ratio of D2 to Cyc mRNA is higher in the medial basal hypothalamus
than in pituitary by both Northern and by RT-PCR analyses (Table 1) in
agreement with our earlier results (12). In both tissues,
hyperthyroidism causes a 2- to 3-fold decrease in the D2 mRNA as
assessed by both techniques (Table 1). Also, the ratio of D2 to
cyclophilin mRNA in GH4C1 cells is 10-fold higher than that in GC
cells. This is consistent with the 8.4-fold higher D2 activity
in GH4C1 than in GC cells as well as the large difference in D2 mRNA
levels by Northern analysis.
Effects of thyroid hormones on D2 expression
Administration of T3 to rats causes a marked decrease
in expression of D2 mRNA in both anterior pituitary and medial basal
hypothalamus (Table 1). However, as assessed either by Northern or by
RT-PCR, the D2 mRNA is not completely suppressed even by T3
administration for 1 week in quantities sufficient to suppress
circulating T4 to undetectable levels. This is consistent
with recent results in intact rats (3, 5, 18). Initial studies showed
that when cells were transferred from normal to thyroid hormone
depleted 10% FBS, there was a 2- to 3-fold increase in the ratio of D2
to cyclophilin mRNA that plateaued at 48–72 h with the increase
slightly greater in GC than GH4C1 cells. This is identical to changes
in D2 activity observed under the same conditions in earlier studies
(27). Exposure of "hypothyroid" GC cells to 100 nM
T3 caused a 50% decrease of the D2 within 2 h similar
to the effect of adding actinomycin D (Fig. 6). The combination of
T3 with actinomycin D caused no faster decrease in D2 mRNA,
indicating that T3 does not alter the half-life of D2 mRNA.
Therefore, T3 rapidly decreases the transcription rate of
the D2 gene. After 4 h T3 exposure, the D2 to
cyclophilin ratio was not decreased further nor did 24 h treatment
of GH4C1 cells cause more than a 40–50% decrease in D2 mRNA (Table 2), similar to the results of prolonged T3 treatment on D2
mRNA in anterior pituitary.
The acute effects of T3 on gene expression can be grouped
into those that are cycloheximide-sensitive, e.g. the
induction of rGH gene transcription, and those that are
cycloheximide-insensitive, such as the increased transcription of D1
mRNA, and both occur in rat pituitary tumor cells (30). The present
studies show for the first time that the T3-induced
suppression of D2 gene transcription falls into the latter category
(Fig. 7) as do the negative effects of T3 on TSH release
from rat pituitary fragments or TSH synthesis in the mouse TtT cell
tumor (31, 32). The maximum T3-induced decrease in D2 mRNA
by T3 in GC or GH4C1 cells over 24 h was 50% (Fig. 7, Table 2), indicating that there is significant
T3-independent transcription of D2 mRNA. This result is in
good agreement with the maximal T3-induced decrease in D2
activity in anterior pituitary and GH4C1 cells in prior studies (27, 33).
D2 activity is also reduced by exposure of pituitary tumor cells to
reverse T3 (Table 2) as shown previously (17, 25, 27). This
iodothyronine is an excellent substrate for D2 with a
Vmax/Km ratio only 2- to 3-fold lower than that
for T4 (3). On the other hand, reverse T3 has a
low affinity for the T3 receptor and does not cause changes
in transcription of T3-dependent genes. In agreement with
this, reverse T3 did not reduce D2 mRNA and, in some
experiments, the D2/cyc ratio was significantly increased (Table 2).
Despite this, reverse T3 caused an 80–90% decrease in D2
activity over 24 h (Table 2). These effects of T3 and
reverse T3 on D2 activity are consistent with the effects
of this iodothyronine by earlier investigators in GH4C1 and GH3 cells
(17, 27). While both of those studies measured D2 activity, a more
recent study in GC cells comparing the time course and dose-response
characteristics concluded that T3 suppressed D2 activity
primarily by a nuclear-receptor-mediated mechanism, whereas the effect
of reverse T3 was posttranslational (25). The fact that
T4 can suppress D2 activity by both mechanisms accounts for
its high potency in reducing D2 activity (16, 18).
The half-life of the D2 message is short, about 2 h. The half-life of the D2 enzyme is approximately 40 min, indicating that the response of D2 to thyroid hormones will be quite rapid whether the decrease in activity is induced by transcriptional or posttranslational effects again consistent with prior studies (17, 18).
Effects of glucocorticoid on D2 mRNA
The effect of glucocorticoids on D2 mRNA are more complex. There
was a doubling of the D2/cyc ratio after 1 h exposure to
10-7 M dexamethasone (Dex), but this change
was transient. The increase was actinomycin-dependent indicating
transcription was required. However, dexamethasone did not effect the
half-life of D2 mRNA (Fig. 9). A combination of T3 and
dexamethasone reduced the DEX effect and, after 2 h, D2 mRNA was
significantly reduced with respect to the zero time control. We are not
aware of previous studies of the effects of glucocorticoid on pituitary
D2 activity, but rat astroglial cells show a 2-fold increase in type 2
deiodinase activity after 2-day exposure to glucocorticoid. In these
cells, glucocorticoid also enhanced the D2 response to 8Br-cAMP or TPA
(34). If glucocorticoid produced a similar effect on the D2 gene in
thyrotrophs, it would contribute to the transient acute suppression of
circulating TSH observed in glucocorticoid-treated rodents and humans
(35, 36, 37).
Effects of cAMP, TRH, and phorbol ester
Although previous studies have shown marked actinomycin
D-dependent increases in D2 activity induced by cAMP derivatives in
astroglial cells and brown adipocytes, the effects in GH4C1 cells are
modest (27, 34, 38, 39), indicating that these are likely to be
cell-type specific. Our results in GC and GH4C1 cells are similar to
those reported by Koenig in that there was only an approximately
40–50% increase in D2/Cyc mRNA ratio in GC cells and a smaller, but
significant, increase by 8-bromo cAMP in GH4C1 cells. Neither TRH nor
TPA caused large increases in D2 mRNA in GH4C1 cells, although Koenig
reported an approximate doubling of 5' D2 activity by these agonists
after a 24-h incubation (Fig. 9B and Table 1) (27). We observed similar
increases (77 and 66%) in D2 activities which thus are larger than
those in D2 mRNA.
In conclusion, we have demonstrated that a carefully controlled semiquantitative PCR technique can be used to explore the mechanism of D2 mRNA regulation in pituitary tumor cell lines. Both T3 and reverse T3 are potent down regulators of D2 activity, but reverse T3 acts exclusively at a posttranslational level. T3 does not alter D2 mRNA half-life but rapidly reduces D2 transcription by a cycloheximide-independent mechanism. cAMP analogs, previously shown to cause marked actinomycin D-dependent increases in D2 activity in astroglial cells or brown adipocytes, have minimal effects on D2 message in pituitary tumor cell lines, suggesting that these responses may be cell-type specific. The effect of T3 in pituitary tumor cells reproduces in magnitude those observed in vivo in both anterior pituitary and in the medial basal hypothalamus. The precision and sensitivity of the method indicates that it can serve as an important tool in further evaluation of the regulation of the D2 gene in small tissue samples.
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Acknowledgments |
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Footnotes |
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Received April 13, 1998.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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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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O. Araki, T. Morimura, T. Ogiwara, H. Mizuma, M. Mori, and M. Murakami Expression of Type 2 Iodothyronine Deiodinase in Corticotropin-Secreting Mouse Pituitary Tumor Cells Is Stimulated by Glucocorticoid and Corticotropin-Releasing Hormone Endocrinology, October 1, 2003; 144(10): 4459 - 4465. [Abstract] [Full Text] [PDF]
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 P. Cettour-Rose, A. G. Burger, C. A. Meier, T. J. Visser, and F. Rohner-Jeanrenaud Central stimulatory effect of leptin on T3 production is mediated by brown adipose tissue type II deiodinase Am J Physiol Endocrinol Metab, November 1, 2002; 283(5): E980 - E987. [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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R. Martinez-deMena, A. Hernandez, and M.-J. Obregon Triiodothyronine is required for the stimulation of type II 5'-deiodinase mRNA in rat brown adipocytes Am J Physiol Endocrinol Metab, May 1, 2002; 282(5): E1119 - E1127. [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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 H. Mizuma, M. Murakami, and M. Mori Thyroid Hormone Activation in Human Vascular Smooth Muscle Cells : Expression of Type II Iodothyronine Deiodinase Circ. Res., February 16, 2001; 88(3): 313 - 318. [Abstract] [Full Text] [PDF]
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 J Yu, M P A Ebert, S Miehlke, H Rost, U Lendeckel, A Leodolter, M Stolte, E Bayerdorffer, and P Malfertheiner alpha -Catenin expression is decreased in human gastric cancers and in the gastric mucosa of first degree relatives Gut, May 1, 2000; 46(5): 639 - 644. [Abstract] [Full Text] [PDF]
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J. Steinsapir, A. C. Bianco, C. Buettner, J. Harney, and P. R. Larsen Substrate-Induced Down-Regulation of Human Type 2 Deiodinase (hD2) Is Mediated through Proteasomal Degradation and Requires Interaction with the Enzyme's Active Center Endocrinology, March 1, 2000; 141(3): 1127 - 1135. [Abstract] [Full Text] [PDF]
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T. Bartha, S.-W. Kim, D. Salvatore, B. Gereben, H. M. Tu, J. W. Harney, P. Rudas, and P. R. Larsen Characterization of the 5'-Flanking and 5'-Untranslated Regions of the Cyclic Adenosine 3',5'-Monophosphate-Responsive Human Type 2 Iodothyronine Deiodinase Gene Endocrinology, January 1, 2000; 141(1): 229 - 237. [Abstract] [Full Text] [PDF]
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 J. R. Gasdaska, J. W. Harney, P. Y. Gasdaska, G. Powis, and M. J. Berry Regulation of Human Thioredoxin Reductase Expression and Activity by 3'-Untranslated Region Selenocysteine Insertion Sequence and mRNA Instability Elements J. Biol. Chem., September 3, 1999; 274(36): 25379 - 25385. [Abstract] [Full Text] [PDF]
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B. Gereben, T. Bartha, H. M. Tu, J. W. Harney, P. Rudas, and P. R. Larsen Cloning and Expression of the Chicken Type 2 Iodothyronine 5'-Deiodinase J. Biol. Chem., May 14, 1999; 274(20): 13768 - 13776. [Abstract] [Full Text] [PDF]
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 Y. I. Choi, S. H. Jeon, J. Jang, S. Han, J. K. Kim, H. Chung, H. W. Lee, H.-Y. Chung, S. D. Park, and R. H. Seong Notch1 confers a resistance to glucocorticoid-induced apoptosis on developing thymocytes by down-regulating SRG3 expression PNAS, August 28, 2001; 98(18): 10267 - 10272. [Abstract] [Full Text] [PDF]
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